UNIVERSITY  OF  CALIFORNIA 
AT   LOS  ANGELES 


METHODS  IN  PLANT  HISTOLOGY 


THE  UNIVERSITY  OP  CHICAGO  PRESS 
CHICAGO,  ILLINOIS 


THE  BAKER  &  TAYLOR  COMPANY 

NEW  YORK 

THE  CAMBRIDGE  UNIVERSITY  PRESS 

LONDON 

THE  MARUZEN-KABUSHIKI-KAISHA 

TOKYO,  OSAKA,  KYOTO,  FTKUOKA,  6ENDAI 

THE  MISSION  BOOK  COMPANY 


METHODS  IN 

PLANT  HISTOLOGY 


CHARLES  J.  CHAMBERLAIN,  A.M.,  PH.D. 

Professor  of  Botany  in  the  University  of  Chicago 


THIRD  REVISED  EDITION 


THE  UNIVERSITY  OF  CHICAGO  PRESS 
CHICAGO,  ILLINOIS 


COPYRIGHT  1901, 1905,  1915,  BY 
THE  UNIVERSITY  OF  CHICAGO 


All  Rights  Reserved 


Published  June  1901 
Second  Edition  October  1905 

Third  Edition  May  1915 

Second  Impression  December  1916 

Third  Impression  July  1920 


Composed  and  Printed  By 

The  University  of  Chicago  Pr 

Chicago,  Illinois.  U.S.A. 


QK 

^H3 

C35  - 


PREFACE  TO  THE  FIRST  EDITION 

This  book  has  grown  out  of  a  course  in  histological  technic 
conducted  by  the  author  at  the  University  of  Chicago.  The  course 
has  also  been  taken  by  non-resident  students  through  the  Extension 
Division  of  the  University.  The  Methods  were  published  over  a 
year  ago  as  a  series  of  articles  in  the  Journal  of  Applied  Microscopy, 
and  have  called  out  numerous  letters  of  commendation,  criticism, 
suggestion,  and  inquiry.  The  work  has  been  thoroughly  revised  and 
enlarged  by  about  one-half.  It  is  hoped  that  the  criticism  and 
suggestion,  and  also  the  experience  gained  by  contact  with  both 
resident  and  non-resident  students,  have  made  the  directions  so 
definite  that  they  may  be  followed,  not  only  by  those  who  work 
in  a  class  under  the  supervision  of  an  instructor,  but  also  by  those 
who  must  work  in  their  own  homes  without  any  such  assistance. 

More  space  has  been  devoted  to  the  paraffin  method  than  to 
any  other,  because  it  has  been  proved  to  be  better  adapted  to  the 
needs  of  the  botanist.  The  celloidin  method,  the  glycerin  method, 
and  freehand  sectioning  are  also  described,  and  their  advantages 
and  disadvantages  are  pointed  out. 

The  first  part  of  the  book  deals  with  the  principles  of  fixing 
and  staining,  and  the  various  other  processes  of  microtechnic, 
while  in  the  later  chapters  these  principles  are  applied  to  specific 
cases.  This  occasions  some  repetition,  but  the  mere  presentation 
of  general  principles  will  not  enable  the  beginner  to  make  good 
mounts. 

The  illustrations  and  notes  in  the  later  chapters  are  not  intended 
to  afford  a  study  of  general  morphology,  but  they  merely  indicate 
to  students  with  a  limited  knowledge  of  plant  structures  the  principal 


PREFACE  TO  THE  THIRD  EDITION 

The  continued  appreciation  accorded  to  Methods  in  Plant  His- 
tology has  exhausted  the  second  edition.  Since  that  edition  appeared, 
methods  have  become  more  and  more  exact,  so  that  the  present 
volume  is  practically  a  new  book.  The  general  arrangement  of  the 
subject-matter,  and  directions  for  collecting  material  and  for  secur- 
ing reproductive  phases  in  the  Algae  and  Fungi  have  been  retained, 
and  a  chapter  on  "Photomicrographs  and  Lantern  Slides"  (chap,  xii) 
has  been  added. 

Great  improvements  have  been  made  in  the  paraffin  method,  so 
that  sections  are  easily  cut  which  were  impossible  ten  years  ago, 
while  ten  years  of  added  experience  with  the  Venetian  turpentine 
method  have  made  it  possible  to  describe  it  so  definitely  that  even 
the  beginner  should  find  no  serious  difficulty. 

The  author  is  deeply  indebted  to  his  colleague,  Dr.  W.  J.  G. 
Land,  for  numerous  suggestions  and  improvements  covering  the 
whole  field  of  microtechnic.  He  is  also  greatly  indebted  to  Dr.  S. 
Yamanouchi  for  many  improvements  in  the  methods  applicable  to 
Algae  and  mitotic  figures. 

Corrections  and  suggestions  will  be  heartily  appreciated. 

CHARLES  J.  CHAMBERLAIN 
CHICAGO 
May,  1915 


viii 


CONTENTS 
PART  I 

PAGE 

INTRODUCTION 3 

CHAPTER  I.    APPARATUS 5 

CHAPTER  II.     REAGENTS 17 

Killing  and  Fixing  Agents 17 

Dehydrating  Agents 31 

Formulae  for  Alcohols 33 

Clearing  Agents 34 

Miscellaneous  Reagents 37 

CHAPTER  III.    STAINS  AND  STAINING 38 

The  Haematoxylins 40 

The  Carmines 49 

TheAnilins 50 

Combination  Stains 59 

CHAPTER  IV.     GENERAL  REMARKS  ON  STAINING 65 

Selection  of  a  Stain 65 

Theories  of  Staining 66 

Practical  Hints  on  Staining 69 

CHAPTER  V.    TEMPORARY  MOUNTS  AND  MICROCHEMICAL  TESTS      .  72 

CHAPTER  VI.    FREEHAND  SECTIONS 80 

CHAPTER  VII.    THE  GLYCERIN  METHOD 92 

CHAPTER  VIII.    THE  VENETIAN  TURPENTINE  METHOD    ....  97 

CHAPTER  IX.    THE  PARAFFIN  METHOD 102 

Killing  and  Fixing 102 

Washing 104 

Hardening  and  Dehydrating 105 

Clearing 106 

The  Transfer  from  Clearing  Agent  to  Paraffin 107 

The  Paraffin  Bath 108 

Imbedding 109 

Cutting Ill 

Fixing  Sections  to  the  Slide 113 

Removal  of  the  Paraffin 115 


x  Methods  in  Plant  Histology 

PAGE 

Removal  of  Xylol  or  Turpentine   . 115 

Transfer  to  the  Stain 116 

Dehydrating 116 

Clearing 116 

Mounting  in  Balsam 117 

A  Tentative  Schedule  for  Paraffin  Sections  . 117 

CHAPTER  X.    THE  CELLOIDIN  METHOD 119 

CHAPTER  XI.    SPECIAL  METHODS 125 

Very  Large  Sections 125 

Stony  Tissues 126 

Petrifactions 126 

Thick  Sections 128 

Land's  Gelatin  Method 128 

Schultze's  Maceration  Method 129 

Protoplasmic  Connections  . 129 

Staining  Cilia • 131 

Mitochondria 132 

Canaliculi 134 

Vascular  Bundles  in  Living  Tissues 135 

Staining  Living  Structures 135 

CHAPTER  XII.    PHOTOMICROGRAPHS  AND  LANTERN  SLIDES    ...  136 

Photomicrographs 136 

Lantern  Slides 141 

PART  II 

SPECIFIC  DIRECTIONS 151 

CHAPTER  XIII.    MYXOMYCETES  AND  SCHIZOPHYTES     .....  152 

CHAPTER  XIV.    CHLOROPHYCEAE 162 

CHAPTER  XV.    PHAEOPHYCEAE 181 

CHAPTER  XVI.    RHODOPHYCEAE .     .     .     .     .  188 

CHAPTER  XVII.    FUNGI .     .     .     ,     ,  r .:     .  192 

Phycomycetes .  192 

Hemiascomycetes 196 

Ascomycetes 196 

Lichens ....,,.  202 

Basidiomycetes .  202 

CHAPTER  XVIII.    BRYOPHYTES— HEPATICAE .207 

CHAPTER  XIX.    BRYOPHYTES — Musci  216 


Contents  xi 

PAGE 

CHAPTER  XX.    PTERIDOPHYTES — LYCOPODIALES 221 

CHAPTER  XXI.    PTERIDOPHYTES— EQUISETALES      .     .     .     ...  227 

CHAPTER  XXII.    PTERIDOPHYTES — OPHIOGLOSSALES 230 

CHAPTER  XXIII.    PTERIDOPHYTES — FILICALES 233 

CHAPTER  XXIV.     SPERMATOPHYTES — GYMNOSPERMS 244 

Cycadales 244 

Ginkgoales 250 

Coniferales 250 

CHAPTER  XXV.    SPERMATOPHYTES — ANGIOSPERMS 261 

CHAPTER  XXVI.    USING  THE  MICROSCOPE 280 

CHAPTER  XXVII.    LABELING  AND  CATALOGING  PREPARATIONS  .      .  288 

CHAPTER  XXVIII.    A  CLASS  LIST  OF  PREPARATIONS       .     .     .     .  290 

CHAPTER  XXIX.    FORMULAE  FOR  REAGENTS 297 

INDEX   .  311 


PART  I 


INTRODUCTION 

The  technic  of  fifty  years  ago,  judged  by  modern  standards,  was 
very  crude;  the  microscopes  of  that  time,  while  no  worse  than  the 
preparations,  could  not  show  the  details  which  interest  investigators 
today.  Many  objects,  like  pollen  grains,  were  examined  without 
sectioning.  The  pollen  grain  of  a  lily,  if  placed  upon  a  dark  back- 
ground, is  barely  visible  to  the  naked  eye;  but  with  modern  methods, 
such  a  pollen  grain  can  be  cut  into  fifty  sections,  the  sections  can  be 
mounted  and  stained  without  getting  them  out  of  order,  a  photo- 
micrograph can  be  made  from  the  preparation  and  a  lantern  slide 
from  the  photomicrograph,  and  finally  there  appears  upon  the  screen 
a  pollen  grain  ten  feet  long,  with  nuclei  a  foot  in  diameter,  nucleoli 
like  baseballs,  and  starch  grains  as  large  as  walnuts.  Impossible  as 
this  may  seem,  such  preparations  are  easily  made,  and  investigators 
are  now  showing  clearly  the  nature  of  structures  which,  only  ten  years 
ago,  were  good  subjects  for  philosophical  botanists,  who  are  happier 
with  preparations  which  leave  more  freedom  for  the  imagination. 

Modern  technic  is  very  complicated,  and  to  the  beginner  the 
numerous  details  may  seem  bewildering,  but  every  detail  must  be 
mastered  if  the  final  mount  is  to  be  worth  anything.  By  following 
the  various  schedules,  even  in  a  slavish  way,  fairly  good  mounts 
have  been  obtained  at  the  first  trial;  but  to  gain  any  independence 
and  to  secure  the  best  results,  the  student  should  understand  the 
reason  for  each  step  in  the  whole  schedule.  Only  then  will  he 
become  able  to  make  such  variations  as  individual  cases  may  require. 
The  horizon  should  broaden  as  the  student  advances,  and  he  should 
see  that  even  such  diverse  methods  as  the  Freehand  Method,  the 
Venetian  Turpentine  Method,  and  the  Paraffin  Method  have 
certain  fundamental  principles  in  common. 

Everyone  who  intends  to  become  an  investigator  should  study 
technic  with  the  intention  of  using  it  in  his  researches.  Many  regard 
the  making  of  mounts  as  mere  mechanical  drudgery  which  can  be 
done  by  an  assistant,  but  such  armchair  investigators  are  likely  to 

3 


4  Methods  in  Plant  Histology 

draw  false  conclusions  or  to  become  scholastic  grafters,  according 
as  the  assistant  is  mediocre  or  talented.  Some  time-honored  theories 
would  have  been  abandoned  long  ago  if  certain  prominent  investi- 
gators had  not  relied  upon  comparatively  untrained  assistants  for 
their  mounts.  Benjamin  Franklin's  advice,  "If  you  would  have 
your  business  done,  go;  if  not,  send,"  applies  very  well  to  the  case 
in  hand. 

Finally,  do  not  imagine  that  you  must  make  an  elaborate  perma- 
nent mount  before  material  is  worth  examining  with  the  microscope. 
Look  at  living  material  whenever  possible;  make  freehand  sections, 
or  tease  with  needles,  and  thus  make  that  preliminary  survey  which 
should  always  precede  the  study  of  permanent  mounts. 


CHAPTER  I 
APPARATUS 

The  amount, of  apparatus  required  for  histological  work  varies, 
temporary  mounts,  glycerin  mounts,  and  freehand  sections  requiring 
only  a  razor  and  a  microscope,  while  the  paraffin  method,  which 
represents  the  highest  development  of  technic,  brings  into  use  nearly 
all  the  equipment  of  the  histological  laboratory.  The  following 
list  includes  only  the  apparatus  necessary  for  making  preparations: 
a  microscope;  a  microtome;  a  razor;  a  hone  and  a  good  razor  strop; 
a  paraffin  bath;  a  turntable;  a  scalpel;  a  pair  of  needles;  a  pair  of 
scissors;  a  pair  of  forceps;  staining-dishes;  solid  watch  glasses; 
bottles;  a  graduate  (50  or  100  c.c.);  pipettes;  slides,  1X3  inches; 
round  covers,  18  mm.  or  f  inch  in  diameter;  and  square  covers,  £  inch. 
Longer  covers  will  be  needed  for  some  of  the  serial  sections. 

A  microscope  should  have  a  rack  and  pinion  coarse  adjustment, 
a  fine  adjustment,  two  eyepieces  magnifying  about  four  and  eight 
diameters,  a  low-power  objective  of  about  16  mm.  focus,  and  a 
high-power  objective  of  about  4  mm.  focus,  a  double  nosepiece,  an 
iris  diaphragm,  and  an  Abbe  condenser.  A  cheap  and  practical 
form  is  shown  in  Fig.  1,  and  similar  instruments  are  for  sale  by  all 
the  leading  companies. 

Since  the  chemicals  used  in  histological  technic  are  likely  to 
damage  the  stage  and  substage  of  the  microscope,  it  is  well  to  place 
upon  the  stage  a  piece  of  glass  three  or  four  inches  square.  A  lantern- 
slide  cover  is  just  right  for  this  purpose.  It  is  not  necessary  to 
fasten  it  to  the  stage,  since  it  is  merely  for  protection  while  examin- 
ing slides  which  are  wet  with  reagents.  In  our  own  laboratory  we 
use  for  examining  wet  slides  a  cheap  microscope  with  only  a  single 
low-power  objective  and  a  single  ocular. 

Some  knowledge  of  the  structure  and  optics  of  the  microscope  is 
necessary  if  one  is  to  use  it  effectively.  Why  are  there  so  many 
diaphragms  ?  Why  is  there  an  arrangement  for  raising  and  lowering 

5 


6 


Methods  in  Plant  Histology 


the  condenser  ?  Why  does  the  mirror  bar  swing  ?  Why  is  one  side 
of  the  mirror  plane  and  the  other  concave?  Everyone  who  uses 
even  a  cheap  microscope  should  know  the  answers  to  questions  like 
these.  All  the  leading  manufacturers  furnish,  free  of  charge,  booklets 

explaining  the  construction 
of  the  microscope  and  giving 
practical  directions  for  its 
care  and  use. 

Aside  from  the  micro- 
scope itself,  the  microtome 
is  the  most  important  piece 
of  apparatus  in  the  labora- 
tory. In  recent  years  there 
has  been  considerable  im- 
provement in  microtomes, 
but  we  still  have  two  gen- 
eral types,  the  sliding  and 
the  rotary. 

The  cheapest  microtomes 
which  have  proved  to  be 
efficient  for  general  work  are 
simple  forms  of  the  sliding 
microtome  like  the  one  shown 
in  Fig.  2.  It  should  be  pro- 
vided with  a  clamp  which 
will  hold  any  kind  of  a  knife 
(Fig.  3).  For  large  or  hard 
objects  the  weakness  of  these 
small  instruments  is  evident 
from  the  figures. 
Where  expense  is  not  too  great  an  objection,  a  larger  microtome 
should  be  secured.  There  is  great  difference  of  opinion  as  to  the 
relative  merits  of  the  sliding  and  rotary  types.  As  far  as  convenience 
and  rapidity  are  concerned,  the  rotary  microtome  is  unquestionably 
superior;  further,  it  will  produce  good  sections  with  less  care  and  skill, 
because  the  movements  are  automatic.  The  fact  that  a  ribbon 


FIG.  1. — An  efficient  microscope  of  moder- 
ate price.  The  leading  optical  companies  put 
the  same  objectives  and  oculars  upon  such  in- 
struments as  upon  their  most  expensive  stands. 


Apparatus 


carrier  is  so  easily  used  with  the  rotary  is  another  great  advantage. 
But  the  sliding  microtome  also  has  its  advantages.     Obviously,  for 


sections  of  stems  and 
general  celloidin  work, 
where  the  knife  is  used 
in  a  very  oblique  posi- 
tion, it  is  not  only 
superior,  but  it  is  the 
only  type  which  has 
proved  to  be  efficient. 
Attempts  to  place  the 
knife  in  an  oblique 
position  in  rotary 
microtomes  have  not 
been  encouraging.  For 
very  thin  paraffin  sec- 
tions the  advantages 
of  the  sliding  micro- 
tome are  such  as  appeal  only  to  the  expert.  With  both  rotary  and 
sliding  types,  a  little  of  the  paraffin  is  sure  to  stick  to  the  side  of  the 


FIG.  3. — Clamp  to  hold  an  ordinary 
razor,  a  heavy  microtome  knife,  or  a 
Stickler's  holder. 


8  Methods  in  Plant  Histology 

knife  next  the  object  after  every  section.  Unless  this  be  wiped  off, 
the  face  of  the  block  is  dragged  across  it  and  the  next  section  is 
damaged  even  before  it  is  cut.  The  side  of  the  knife  next  the  object 
should  be  wiped  with  the  finger,  theoretically  after  every  section. 
It  is  very  inconvenient  to  wipe  the  knife  in  a  rotary  microtome. 
Another  advantage  of  the  sliding  type  is  easy  to  feel  but  difficult  to 
describe:  in  the  rotary  microtome  the  stroke  is  so  automatic  that 
there  is  little  room  for  skill,  but  in  the  sliding  microtome,  with  one's 
hand  on  the  sliding  block,  little  variations  in  the  stroke,  variations 
which  become  instinctive,  give  the  expert  a  control  not  yet  attained 
in  the  rotary  forms. 

Amateurs,  and  even  professional  botanists  who  have  little 
aptitude  in  the  use  of  machines,  had  better  rely  upon  the  rotary 
microtome.  However,  no  better  comment  on  the  comparative  merits 
of  the  two  forms  could  be  given  than  the  practice  of  an  expert 
technician  in  our  own  laboratory,  who  uses  a  rotary  microtome  when 
making  sections  for  ordinary  class  work,  but  who  turns  to  a  sliding 
microtome  of  the  Jung-Thoma  pattern  when  cutting  sections  for 
his  own  research. 

Attempts  have  been  made  to  combine  the  advantages  of  the 
sliding  and  rotary  types.  The  "Precision  Microtome,"  made  by 
Bausch  &  Lomb,  has  found  favor  in  some  circles.  It  has  the  sliding 
movement,  allows  an  oblique  position  of  the  knife,  and  is  operated 
like  a  rotary  microtome.  Recently,  a  much-improved  microtome, 
made  by  the  Spencer  Lens  Co.,  has  been  winning  favor.  It  is  a 
rotary,  but  even  surpasses  the  sliding  microtome  in  precision  and 
stability.  Its  effectiveness  depends,  in  large  measure,  upon  its 
simple  but  rigid  clamp  for  holding  the  object.  This  microtome, 
fitted  with  Dr.  Land's  apparatus  for  cooling  both  the  knife  and  the 
paraffin  block,  is  shown  in  Fig.  4.  Streams  of  ice  water  flow  under 
the  knife  and  through  the  hollow  block  to  which  the  paraffin  is 
fastened.  From  paraffin  with  a  melting-point  of  52°  C.,  or  even 
somewhat  less,  uniform  ribbons  1  /x  in  thickness  can  be  secured. 
If  material  has  been  imbedded  in  paraffin  of  52°  C.  and  it  should 
be  desirable  to  cut  sections  at  15  /*  to  20  M,  warm  water  can  be 
used. 


Apparatus 


9 


A  motor,  as  shown  in  the  figure,  not  only  produces  a  very  even 
stroke,  but  leaves  both  hands  free  to  take  care  of  the  ribbon. 

Microtome  knives  are  available  everywhere  and,  when  perfectly 
sharpened,  are  unsurpassed.  Those  who  sharpen  knives  for  sur- 
geons can  grind  out  nicks,  but  they  do  not  know  how  to  sharpen  a 
microtome  knife  and  they  cannot  be  taught;  they  sharpen  knives  for 
Dr.  Carver  and  Dr.  Cutterout. 


FIG.  4. — Spencer  rotary  microtome  fitted  with  Land's  apparatus  for  temperature 
control,  as  described  in  the  Botanical  Gazette,  June,  1914. 

In  recent  years  several  clamps  have  been  devised  to  hold  the 
blade  of  the  Gillette  safety  razor,  the  hard,  even  edge  of  which  is 
very  satisfactory  for  microtome  sections.  After  dealers  had  ignored 
our  suggestions,  Mr.  A.  W.  Strickler,  at  our  request,  devised  the 
form  of  holder1  shown  in  Fig.  5.  It  is  made  of  brass  and  can  be 
used  in  either  rotary  or  sliding  microtomes.  The  sectional  view 
shows  that  the  two  pieces  of  the  holder  are  curved,  a  feature  which 
insures  great  rigidity.  It  is  neither  necessary  nor  desirable  to  have 

» This  holder  may  be  obtained  from  Mr.  A.  W.  Strickler,  5654  Kenwood  Avenue, 
Chicago,  Illinois,  for  $3.00,  postpaid  $3. 15. 


10 


Methods  in  Plant  Histology 


pins  fitting  the  three  holes  in  the  blade,  since  they  add  nothing  to 
the  rigidity  and  even  interfere  with  the  insertion  and  adjustment  of 
the  knife.  The  knife  should  not  project  more  than  a  millimeter 
beyond  the  holder.  With  the  Gillette  blade  in  this  holder,  we  have 


r 


FIG.  5.— Stickler's  clamp  for  holding  Gillette  blades 

cut  smooth  sections,  2  and  3/t  in  thickness,  and  have  cut  large 
sections  2  cm.  in  diameter  and  15  /z  in  thickness,  even  such  refractory 
objects  as  the  strobili  of  Isoetes  and  Selaginella  cutting  as  smoothly 
as  with  a  first-class  microtome  knife.  When  the  success  of  the 


Fio.  6.— Scalpels  made  from  Gillette  blades,  showing  a  blade  which  has  been  cut 
into  pieces  with  shears,  three  of  the  pieces  soldered  to  nails  with  flattened  heads,  and  a 
scalpel  used  in  an  ordinary  needle-holder. 

holder — or  rather,  its  sale — became  evident,  two  prominent  optical 
companies,  without  any  apologies  or  reference  to  Mr.  Strickler, 
began  to  manufacture  it  and  advertised  it  in  their  catalogs.  Up  to 
this  time  their  holders  are  much  inferior  to  Mr.  Strickler's,  doubtless 
because  they  overlooked  a  very  important,  but  very  obscure  detail. 


Apparatus  11 

When  the  Gillette  blade  begins  to  lose  a  little  of  its  effectiveness 
for  microtome  work  it  will  make  two  or  three  scalpels.  With  a  pair 
of  stout  shears,  cut  the  blade  into  pieces,  as  indicated  in  Fig.  6. 
Take  a  small  steel  nail  and  flatten  the  head  and  upper  part  by  laying 
it  upon  a  piece  of  iron  and  hitting  it  with  a  hammer,  or  by  squeezing 
it  in  a  vise;  then  solder  the  blade  to  the  nail,  and  use  the  scalpel  in  an 
ordinary  needle  holder,  or  drive  the  nail  into  any  wooden  holder.  A 
dozen  of  these  scalpels  can  be  made  in  ten  minutes. 

The  stout  razors  our  grandfathers  used  to  shave  with  are  excellent 
for  freehand  sectioning  and  even  for  cutting  sections  on  the  micro- 
tome. The  blade  should  be  flat  on  one 
side  (Fig.  7,  A).  Modern  razors  (Fig.  7,  5) 
with  delicate  blades  ground  hollow  on  both 
sides,  are  worthless  for  cutting  sections  of 
plants. 

There  should  be  two  good  hones  :   a  fine 

,  ,  f          ,,  r      •  FIG.  7.  —  The  type  shown 

carborundum    hone    for    the    preliminary    in  A  is  good  for 


sharpening,  and  a  yellow  Belgian  hone  for    work;  that  shown  in  B  is 

worthless  for  microtome  work, 
finishing.      About    10X2f    inches   IS   a   good      but  can  be  used  for  freehand 

size.     If  the  second  hone  be  quite  hard    sections  of  leaves. 
and  the  finishing  skilfully  done,  little   or  no  stropping  may  be 
necessary.     The  best  strops  used  by  barbers  are  satisfactory  for 
microtome  knives. 

There  are  numerous  forms  of  the  paraffin  bath.  '  Those  with  a 
water-jacket,  a  thermometer,  and  a  thermostat  to  maintain  an  even 
temperature  are  the  most  convenient. 

Where  electricity  is  available,  the  electric  thermostat  devised  by 
Dr.  Land  is  immensely  superior.  It  is  clean  and  easily  regulated  and 
maintains  a  more  even  temperature  than  has  yet  been  secured  by 
means  of  gas  regulators.  The  appearance  and  principal  features 
are  shown  in  Figs.  8  and  9.  A  detailed  description  of  this  thermostat 
is  given  in  the  Botanical  Gazette  for  November,  1911.  One  familiar 
with  tools  and  electricity  could  make  this  thermostat  at  an  outlay 
of  about  $3.75.  Mr.  A.  W.  Strickler,  5654  Kenwood  Avenue, 
Chicago,  makes  the  apparatus  complete  and  ready  to  attach  to  the 
bath  for  $15. 


12 


Methods  in  Plant  Histology 


A  bath  which,  if  carefully  watched,  gives  the  very  best  results 
can  be  made  by  any  tinner,  and  is  very  inexpensive.  The  figure 
on  p.  14  shows  the  form  and  dimensions  (Fig.  10).  It  is  made 
of  copper  2  or  3  mm.  in  thickness.  Several  triangular  pieces  may  be 
cut  from  a  single  plate  of  copper.  Brass  may  be  used  instead.  The 
three  legs  should  be  screwed  into  the  triangular  plate.  There  should 
be  two  boxes  to  contain  the  paraffin.  They  should  be  about  10  cm. 
long,  2  cm.  wide,  and  2  cm.  high,  and  should  have  loosely  fitting 


FIG.  8. — Land's  electrical  constant  apparatus,  showing  diagram  of  the  automatic 
switch,  as  described  in  the  Botanical  Gazette,  November,  1911. 

covers.  The  long  box  makes  it  possible  to  have  melted  paraffin  at 
one  end  and  paraffin  just  below  the  melting-point  at  the  other  end. 
By  careful  watching  this  bath  will  give  as  good  results  as  the  most 
expensive  bath  with  its  elaborate  thermostat. 

With  the  disappearance  of  the  glycerin  method,  the  turntable  is 
disappearing  from  the  botanical  laboratory;  but  some  objects,  like 
Nemalion  and  moss  protonema,  are  still  mounted  in  glycerin,  and 
so  one  still  finds  occasional  use  for  this  once  necessary  apparatus.  A 
serviceable  form  is  shown  in  Fig.  11  (p.  14).  The  more  expensive 
turntables  with  devices  for  automatic  centering  present  no  practical 
advantages  and  the  centering  devices  are  often  in  the  way. 


Apparatus  13 

Scalpels  made  from  thin  razor  blades  have  been  mentioned 
already.  For  trimming  paraffin  blocks  and  handling  paraffin  ribbons 
a  more  rigid  scalpel  is  necessary. 

Needles  are  used  so  constantly  that  it  is  well  to  have  clamping 
holders.  However,  if  it  were  not  for  the  trouble  of  inserting  and 


FIG.  9. — Thermostat,  heater,  and  switch  of  Land's  electrical  constant  apparatus 

pulling  out  needles,  nothing  is  quite  equal  to  a  rather  large  handle 
whittled  out  from  a  piece  of  light  pine. 

Scissors  are  seldom  used  in  the  botanical  laboratory  except  for 
cutting  out  labels.  Rather  stout  scissors,  with  blades  about  2 \  inches 
in  length,  are  best  for  general  purposes. 


14 


Methods  in  Plant  Histology 


It  is  convenient  to  have  two  pairs  of  forceps,  a  strong  pair  for 
handling  slides  and  a  delicate  pair  for  handling  covers.  If  there  is 
to  be  only  one  pair,  they  must  be  strong  enough  for  the  slides  but 


FIG.  10. — Paraffin  bath 


not  too  clumsy  for  covers.  Curved  forceps  are  not  necessary;  the 
cover-glass  forceps,  used  by  bacteriologists  in  staining  on  the  cover, 
are  of  no  use  in  botanical  work. 


FIG.   11. — Turntable 

Stender  dishes  are  very  generally  used  in  staining  on  the  slide. 
The  form  shown  in  Fig.  12,  A,  about  60X90  mm.,  is  in  quite  general 
use.  Some  prefer  the  Coplin  jar,  shown  in  Fig.  12,  B.  The  latter 
is  more  troublesome  to  clean,  but  requires  less  of  the  reagent.  Many 
other  forms  are  on  the  market.  When  large  numbers  of  slides  of  the 
same  kind  are  to  be  stained  at  one  time,  small  battery  jars,  holding 
about  a  liter,  may  be  used.  In  this  case,  it  is  well  to  have  a  rack, 


Apparatus 


15 


holding  20  to  30  slides,  so  that  all  may  be  transferred  at  the  same 
time  from  one  reagent  to  another.  With  this  convenience,  it  is  not 
necessary  to  handle  the  slides  separately,  except  at  critical  stages. 

Solid  watch  glasses,  or  Minots,  as  they  are  often  called,  are  always 
useful.  Each  student  should  have  a  dozen  or  more. 

Each  student  should  have  three  bottles  of  about  one  liter  capacity 
for  90  per  cent  alcohol,  absolute  alcohol,  and  xylol.  In  addition, 
half  a  dozen  bottles,  holding  about  100  c.c.,  will  be  useful.  There 
should  be  two  bottles,  holding  about  50  c.c.,  for  clove  oil.  If  one  is 


A  B 

FIG.  12. — Staining-dishes:    A,  Stender  dish;    B,  Coplin  jar 

doing  much  research  work,  it  will  be  convenient  to  have  many  more 
bottles  for  graded  series  of  alcohols  and  xylols. 

There  should  be  a  graduate,  preferably  50  c.c.  or  100  c.c.  If 
the  bottles  are  of  uniform  sizes,  50  c.c.,  100  c.c.,  500  c.c.,  and  1,000'c.c., 
the  student  should  soon  be  able  to  estimate  with  sufficient  accuracy 
for  making  up  reagents  which  do  not  require  extreme  accuracy. 

Three  or  four  pipettes,  or  medicine-droppers,  will  be  useful. 
Occasionally,  the  glass  of  an  ordinary  pipette  thrust  into  a  small 
camera  bulb  will  save  time  in  drawing  off  reagents. 

Slides  and  covers  are  a  constant  expense.  Many  slides  now 
upon  the  market  are  imperfect.  Beware  of  slides  which  are  not 
perfectly  flat.  Be  skeptical  in  regard  to  any  claim  that  slides  are 
already  clean  enough  to  use.  Of  course,  there  should  be  no  bubbles. 
"White"  slides  are  to  be  preferred  to  those  which  appear  greenish  in 


16  Methods  in  Plant  Histology 

the  box.  For  ordinary  class  work,  slides  of  medium  thickness  are 
more  serviceable,  but  for  critical  cytological  work  many  investigators 
prefer  very  thin  slides. 

There  is  never  any  objection  to  very  thin  covers,  except  that  they 
require  care  in  cleaning.  For  mounts  which  are  to  be  used  with  an 
immersion  lens,  it  is  better  to  have  the  cover  of  the  same  width  as 
the  slide.  The  advantage  is  evident,  since  there  is  no  danger  of 
getting  balsam  on  the  cover  when  wiping  off  the  immersion  fluid; 
besides,  one  can  put  sections  to  the  very  edge  of  the  slide  and  still 
be  sure  that  they  will  be  covered.  Since  most  mounts  for  research 
work  are  mounted  under  long  covers  and  are  intended  for  examina- 
tion with  immersion  lenses,  we  should  recommend  covers  of  25X50 
mm.,  or  even  25X60  mm.  Round  covers  are  desirable  only  when 
mounts  are  to  be  sealed  on  a  turntable.  Larger  slides  and  corre- 
spondingly larger  covers  are  needed  for  special  purposes. 

By  consulting  a  catalog,  which  will  be  furnished  by  any  dealer, 
the  beginner  can  determine  what  he  needs  to  buy,  and  what  he  can 
find  substitutes  for,  if  it  is  necessary  to  be  very  economical. 


CHAPTER  II 
REAGENTS 

While  very  few  new  reagents  have  come  into  general  use  since 
the  second  edition  of  this  book  was  published  in  1905,  there  have 
nevertheless  been  important  improvements  in  the  use  of  some  of  the 
time-honored  combinations.  Doubtless  the  number  of  reagents  used 
in  the  histological  laboratory  will  continue  to  increase,  but  the 
improvements  of  the  past  eight  years  have  been  due  more  to  increased 
care  and  precision  in  the  use  of  well-known  reagents  than  to  the  dis- 
covery of  new  combinations  or  ingredients.  The  following  account 
includes  those  which  are  used  constantly,  and  also  a  few  which  are 
used  occasionally.  The  Microtomist's  Vade-Mecum,  by  Lee,  is  written 
from  the  standpoint  of  the  zoologist,  but  it  contains  very  complete 
formulae  for  stains  and  other  reagents. 

A  list  of  reagents,  with  the  quantities  used  by  the  average  student 
in  a  three  months'  course  in  methods,  is  given  in  chap.  xxix. 
"Stains  and  Staining"  are  described  in  chap.  iii. 

KILLING  AND  FIXING  AGENTS 

Usually  the  same  reagent  is  used  for  both  killing  and  fixing. 
The  purpose  of  a  killing  agent  is  to  bring  the  life-processes  to  a 
sudden  termination,  while  a  fixing  agent  is  used  to  fix  the  cells  and 
their  contents  in  as  nearly  the  living  condition  as  possible.  The 
fixing  consists  in  so  hardening  the  material  that  the  various  elements 
may  retain  their  natural  condition  during  all  the  processes  which 
are  to  follow.  Zoologists  often  use  chloroform  or  ether  for  killing  an 
organism,  and  then  use  various  fixing  agents  for  various  tissues.  No 
promptings  of  humanity  restrain  the  botanist  from  the  vivisection  of 
plants,  but  separate  reagents  for  killing  and  fixing  are  sometimes 
used,  e.g.,  material  may  be  killed  by  placing  it  for  a  short  time  in 
Flemming's  fluid,  which  is  a  very  rapid  killing  agent,  after  which  the 
fixing  may  be  completed  in  a  chromo-acetic  solution,  without  any 

17 


18  Methods  in  Plant  Histology 

osmic  acid,  thus  securing  the  advantages  of  a  very  rapid  killing 
without  the  blackening  which  results  from  a  prolonged  treatment 
with  a  solution  containing  osmic  acid. 

Probably  no  process  in  microtechnic  is  in  more  urgent  need  of 
improvement  than  this  first  step  of  killing  and  fixing.  Nearly  all 
of  our  formulae  are  merely  empirical,  for  very  few  botanists  are 
expert  chemists,  and  those  who  have  the  requisite  knowledge  of 
chemistry  are  interested  in  physiological  problems  rather  than  in 
microtechnic.  The  principal  ingredients  of  the  usual  killing  and 
fixing  agents  are:  alcohol,  chloroform,  chromic  acid,  acetic  acid, 
osmic  acid,  formic  acid,  picric  acid,  sulphuric  acid,  platinum  chloride, 
iridium  chloride,  corrosive  sublimate,  and  formalin.  We  shall  con- 
sider first: 

THE  ALCOHOLS 

a)  Ninety-five  Per  Cent  Alcohol. — This  is  in  quite  general  use 
for  material  which  is  needed  only  for  rough  work.  It  is  extremely 
convenient,  since  it  kills,  fixes,  and  preserves  at  the  same  time  and 
needs  no  changing  or  washing.  It  really  has  nothing  to  recommend 
it  for  fine  work.  It  causes  protoplasm  to  shrink,  but  cell  walls 
usually  retain  their  position,  so  that  95  per  cent  alcohol  will  do  for 
freehand  sections  of  wood  and  many  herbaceous  stems;  but  even 
freehand  sections  of  tender  stems,  like  geraniums  and  begonias,  will 
look  better  if  better  reagents  are  employed.  Alcohols  weaker  than 
95  per  cent  are  not  to  be  recommended  as  fixing  agents,  although  70 
per  cent  alcohol,  or  even  50  per  cent,  will  preserve  material  for  habit 
work.  The  time  required  for  fixing  in  95  per  cent  alcohol  is  about 
the  same  as  for  absolute  alcohol.  The  subsequent  treatment  is  the 
same,  except  that  material  to  be  imbedded  in  paraffin  or  celloidin 
must  be  dehydrated  in  absolute  alcohol.  Material  preserved  in 
weaker  alcohols  and  intended  only  for  habit  study  may  be  kept  in  the 
the  reagent  until  needed  for  use.  Unless  some  glycerin  be  added, 
material  left  in  95  per  cent  alcohol  becomes  very  brittle.  Stems, 
roots,  and  similar  objects  may  be  kept  indefinitely  in  a  mixture  of 
equal  parts  of  95  per  cent  alcohol  and  glycerin. 

Methyl  alcohol,  or  wood  alcohol  as  it  is  commonly  called,  serves 
equally  well. 


Reagents  19 

6)  Absolute  (100  Per  Cent)  Alcohol. — This  is  a  fair  killing  and 
fixing  agent,  it  causes  but  little  shrinking  of  the  protoplasm,  and  is 
a  time-saver  if  material  is  to  be  imbedded  in  paraffin.  The  time 
required  for  fixing  in  alcohol  is  very  short.  For  small  fungi,  like 
Eurotium,  1  minute  is  long  enough.  Root-tips  of  the  onion,  anthers 
of  the  lily,  and  similar  objects  require  15  to  30  minutes.  Larger 
objects  may  require  an  hour.  No  washing  is  necessary,  but  all 
plant  tissues  contain  water;  consequently,  if  material  is  to  be 
imbedded  in  paraffin,  the  alcohol  used  for  fixing  should  be  poured 
off  and  fresh  alcohol  added  before  proceeding  with  the  clearing.  If 
material  is  to  be  mounted  in  Venetian  turpentine,  as  is  likely  to  be  the 
case  in  small  filamentous  fungi,  the  transfer  to  the  stain  may  be 
made  directly  from  the  absolute  alcohol.  This  is  only  for  very  small 
forms,  like  Aspergillus;  neither  the  fixing  nor  the  rude  transfer  would 
be  at  all  satisfactory  with  a  form  so  large  as  Saprolegnia. 

Acetic  acid  is  used  with  alcohols  to  counteract  the  tendency  to 
shrink.  One  of  the  most  widely  known  of  the  alcohol  combinations  is 

c)  Carnoy's  Fluid.-— 

Absolute  alcohol 6  parts 

Chloroform 3  parts 

Glacial  acetic  acid 1  part 

The  penetration  of  the  reagent  is  very  rapid.  An  object  like  an 
onion  root-tip  is  doubtless  killed  in  less  than  a  minute,  and  10  to  15 
minutes  is  long  enough  for  fixing  an  object  of  this  size.  Wash  in 
absolute  alcohol,  changing  frequently,  until  there  is  little  or  no  odor 
of  acetic  acid.  For  a  root-tip,  the  entire  process  of  fixing  and  wash- 
ing should  not  require  more  than  an  hour.  It  is  better  to  imbed  in 
paraffin  at  once,  but  when  this  is  not  convenient  the  material  may  be 
transferred  to  85  per  cent  alcohol,  where  it  may  be  left  until  needed. 
Cyanin  and  erythrosin,  fuchsin  and  iodine  green,  and  similar  com- 
binations give  particularly  brilliant  staining  after  this  reagent. 

d)  Acetic  Alcohol. — Farmer  and  Shove  recommend  for  fixing 
root-tips  of  Tradescantia  virginica  a  mixture  of  two  parts  absolute 
alcohol  and  one  part  of  glacial  acetic  acid.     The  mixture  is  allowed 
to  act  for  15  to  20  minutes,  after  which  the  acid  is  washed  out  with 
absolute  alcohol  and  the  material  is  imbedded  as  soon  as  possible. 


20  Methods  in  Plant  Histology 

e)  Formalin  Alcohol. — The  most  satisfactory  of  the  alcohol 
combinations  is  formalin  alcohol.  Various  proportions  are  used  by 
different  workers.  Professor  Lynds  Jones,  who  first  brought  this 
combination  to  my  notice,  added  2  c.c.  of  commercial  formalin  to 
100  c.c.  of  70  per  cent  alcohol.  We  have  used  a  larger  proportion  of 
formalin,  often  as  much  as  6  c.c.  to  100  c.c.  of  70  per  cent  alcohol. 
Results  which  seem  equally  good  have  been  secured  by  adding  4  to 
6  c.c.  of  formalin  to  100  c.c.  of  50  per  cent  alcohol. 

This  fixing  agent  is  ideally  convenient,  for  material  is  simply 
placed  in  it  and  left  until  wanted  for  use,  an  advantage  which  can  be 
fully  appreciated  only  by  those  who  make  extensive  collecting 
expeditions  at  great  distances  from  the  conveniences  of  the  labora- 
tory. For  anatomical  work,  this  combination  is  unsurpassed,  and 
even  for  delicate  material,  like  liverworts  and  fern  prothallia,  the 
results  are  satisfactory.  The  fixing  is  rapid,  as  is  shown  by  the 
abundance  of  mitotic  figures.  However,  for  a  critical  study  of 
mitotic  figures,  other  fixing  agents  are  preferable. 

THE  CHROMIC-ACID  GROUP 

Chromic  acid,  or  solutions  with  chromic  acid  as  a  foundation,  are 
the  most  generally  useful  killing  and  fixing  agents  yet  known  to  the 
botanist.  A  1  per  cent  solution  of  chromic  acid  in  water  gives  good 
results,  but  it  is  better  to  use  the  chromic  in  connection  with  other 
ingredients,  such  as  acetic  acid,  formic  acid,  osmic  acid,  etc.  Chromic 
acid  does  not  penetrate  well,  and  this  is  one  reason  why  it  is  seldom 
used  alone.  Unfortunately  it  precipitates  some  liquid  albuminoids 
in  the  form  of  filaments  and  networks,  which  may  be  mistaken  for 
structural  elements.  In  botanical  work,  acetic  acid  is  nearly  always 
mixed  with  chromic  acid.  The  pickles  of  the  dinner  table  show  that 
acetic  acid  is  a  good  preservative,  and  that  it  causes  little  or  no 
shrinking.  It  penetrates  rapidly,  and  is  likely  to  cause  swelling 
rather  than  shrinking,  thus  counteracting  the  tendency  of  chromic 
acid  to  cause  plasmolysis.  The  swelling  is  as  bad  as  shrinking. 
Solutions  containing  more  than  1  per  cent  of  acetic  acid  are  to  be 
regarded  with  suspicion.  However,  if  the  purpose  is  to  show  the 
topography  of  structures  like  the  egg  and  synergids  in  the  embryo- 


Reagents  21 

4 

sac  of  an  angiosperm,  or  the  free  nuclear  stage  in  the  endosperm 
of  a  gymnosperm,  2  per  cent,  or  even  3  per  cent,  may  be  used;  but 
the  finer  details  of  the  nucleus  and  cytoplasm  are  damaged  by  such 
strong  solutions. 

It  will  be  found  convenient  to  have  in  the  laboratory  the  following 
stock  solution  of  chromo-acetic  acid  from  which  various  solutions 
can  be  made  as  they  are  needed : 

Chromic-acid  crystals 10  g. 

Glacial  acetic  acid 10  c.c. 

Water 1,000  c.c. 

To  make  a  solution  containing  0 . 5  g.  of  chromic  acid  and  2  c.c.  of 
glacial  acetic  acid  to  100  c.c.  of  water,  add  50  c.c.  of  water  to  50  c.c. 
of  the  stock  solution,  and  then  add  to  the  weakened  solution  1 . 5  c.c. 
of  glacial  acetic  acid.  Any  desired  proportions  can  be  secured  in  a 
similar  way.  Weighing  the  crystals  for  every  new  proportion  is 
more  tedious.  The  proportions  of  the  various  ingredients,  for  the 
present  at  least,  must  be  determined  by  experiment.  With  favorable 
objects  like  fern  prothallia,  Spirogyra,  and  other  things  which  can 
be  watched  while  the  fixing  is  taking  place,  suitable  proportions  are 
rather  easily  determined,  because  specimens,  after  being  placed  in 
the  reagent,  may  be  examined  at  frequent  intervals,  and  combinations 
which  cause  plasmolysis  may  be  rejected  and  different  proportions 
tried  until  satisfactory  results  are  secured.  For  example,  fern  pro- 
thallia might  be  placed  in  the  following  solution:  chromic  acid,  2  g.; 
acetic  acid,  1  c.c.;  and  water,  97  c.c.  If  plasmolysis  takes  place, 
weaken  the  chromic  or  strengthen  the  acetic,  since  the  chromic 
has  a  tendency  to  produce  contraction,  and  the  acetic  to  cause 
swelling.  A  good  fixing  agent  for  fern  prothallia  can  be  made  by 
adding  50  c.c.  of  water  and  1  c.c.  of  glacial  acetic  acid  to  50  c.c.  of 
the  stock  solution.  This  solution  will  cause  practically  no  plas- 
molysis, and  the  fixing  is  thorough,  but  it  must  be  remembered 
that  the  proportion  of  acetic  acid  is  rather  high  for  cytological 
details.  A  combination  may  be  quite  satisfactory  for  fern  pro- 
thallia and  still  fail  to  give  good  results  with  Spirogyra,  and  a  com- 
bination which  succeeds  very  well  with  Spirogyra  may  not  succeed 
at  all  with  Vaucheria.  For  very  critical  work  the  most  favorable 


22  Methods  in  Plant  Histology 

0 

proportions  must  be  determined  for  the  particular  plant  under 
investigation.  In  observing  the  effect  of  the  fixing  one  can  determine 
whether  there  is  any  noticeable  plasmolysis  or  distortion,  but  whether 
the  fixing  is  thorough  can  be  determined  only  by  noting  how  the 
tissues  endure  the  subsequent  processes.  When  the  effect  of  the 
reagent  cannot  be  observed  directly,  it  is  well  to  make  a  freehand 
section  and  thus  determine  whether  plasmolysis  takes  place.  It 
is  not  safe  to  judge  the  action  of  a  fixing  agent  by  the  appearance 
of  sections  cut  from  material  which  has  been  imbedded  in  paraffin, 
because  shrinking  of  the  cell  contents  often  takes  place  during  the 
transfer  from  absolute  alcohol  to  the  clearing  agent  or  during  infil- 
tration with  paraffin,  and  sometimes  even  during  later  processes. 
When  there  is  doubt  as  to  proportions,  we  should  suggest  2  c.c. 
chromic  acid,  3  c.c.  acetic  acid,  and  300  c.c.  water  as  a  good  formula 
for  most  purposes. 

A  large  quantity  of  the  fixing  agent  is  required  and  it  cannot  be 
used  again.  The  volume  of  the  fixing  agent  should  be  at  least  25 
times  that  of  the  material  to  be  fixed.  We  use  about  50  volumes  of 
the  fixing  agent  to  one  of  the  material. 

The  time  required  for  fixing  undoubtedly  varies  with  different 
objects,  but  even  a  delicate  object,  like  Spirogyra,  which  is  pene- 
trated immediately,  should  remain  in  the  fixing  fluid  for  18  to  24 
hours.  Most  botanists  leave  material  like  onion  root-tips  and  lily 
ovaries  in  the  chromo-acetic  acid  about  24  hours.  Some  recommend 
longer  periods.  Christman,  in  his  work  on  rusts,  left,  material  for 
three  days  in  Flemming's  fluid,  a  much  more  vigorous  agent  than 
the  chromo-acetic  acid:  We  have  often  imbedded  material  which 
had  been  in  chromo-acetic  acid  for  several  days,  and  it  seemed  to 
have  suffered  no  injury.  It  is  well  known  that  zoologists  allow 
fixing  agents  like  Miiller's  fluid  and  Erlicki's  fluid  to  act  for  weeks 
before  the  material  is  passed  on  to  the  next  stage,  and  it  may  well  be 
questioned  whether  botanists  have  not  made  a  mistake  in  allowing 
the  chromic  solutions  to  act  for  so  short  a  time.  More  rapid  pene- 
tration, and  consequently  more  immediate  killing,  can  be  secured  if 
the  reagent  is  kept  warm  (30°  to  40°  C.)«  The  warming  also  shortens 
the  time  required  for  fixing,  but,  for  cytological  work,  it  is  quite 


Reagents  23 

possible  that  the  danger  of  producing  artifacts  may  be  increased  by 
the  heat. 

After  fixing  is  complete,  all  reagents  containing  chromic  acid 
as  an  ingredient  should  be  washed  out  with  water.  Running  water 
is  desirable,  and  where  this  is  not  convenient  the  water  must  be 
changed  frequently. 

About  8  or  10  hours  should  be  long  enough  for  filamentous  algae 
and  fungi,  which  are  immediately  penetrated  by  the  water.  It  is  a 
good  plan  to  start  the  washing  in  the  morning  and  let  the  material 
wash  all  day.  For  fern  prothallia,  onion  root-tips,  lily  anthers,  and 
any  material  from  such  a  size  up  to  cubes  a  centimeter  square,  let  the 
material  wash  for  24  hours.  Even  for  delicate  algae,  24  hours  does 
no  damage,  and  some  of  the  best  cytologists  prefer  the  prolonged 
washing. 

Many  methods  have  been  devised  for  insuring  thorough  washing 
and  for  facilitating  the  process.  The  most  obvious  method  is  to 
allow  a  gentle  stream  of  water  to  flow  into  the  Stender  dish  or  bottle 
containing  the  material.  There  is  little  danger  in  this  method  if  the 
material  is  heavy  enough  to  remain  at  the  bottom:  the  only  objection 
is  that  much  of  the  water  never  reaches  the  bottom  where  it  is  needed. 
If  material  is  lighter,  tie  a  piece  of  cheese-cloth  over  the  mouth  of  the 
bottle. 

An  apparatus  for  washing  several  collections  at  one  time  may  be 
made  as  follows :  Get  a  piece  of  f -inch  lead  pipe,  bore  holes  about  y5^ 
or  f  inch  in  diameter  and  about  1|  inches  apart,  put  a  short  rubber 
tube  in  each  hole  and  the  glass  part  of  a  pipette  in  the  end  of  each 
rubber  tube.  Connect  the  lead  tube  with  the  faucet  by  a  large 
rubber  tube.  A  still  better  way  is  to  bore  T\-inch  holes  in  the  lead 
tube,  screw  into  these  holes  short  brass  tubes,  and  then  fasten  the 
pipettes  to  the  brass  tubes  with  thin  rubber  tubes. 

If  there  are  no  facilities  for  working  with  metal,  take  a  wooden 
box  about  6  inches  wide,  18  inches  long,  and  4  inches  deep;  bore 
f-inch  holes  in  the  bottom,  and  into  each  hole  put  a  piece  of  rubber 
tubing  about  4  or  5  inches  in  length.  The  pipettes  can  be  fastened 
in  the  ends  of  these  rubber  tubes.  Place  the  box  under  the  tap.  In 
the  botanical  laboratory  at  Woods  Hole,  Massachusetts,  large 


24  Methods  in  Plant  Histology 

quantities  of  material  are  washed  at  one  time  by  using  an  ordinary 
washtub  with  the  bottom  arranged  as  just  described  for  the  box. 
If  one  is  using  such  a  large  box  or  tub  and  does  not  need  all  the 
streams  of  water,  the  tubes  not  in  use  may  be  closed  by  means  of 
clamps. 

The  following  is  a  simple  and  effective  method :  Cut  f -  or  j-inch 
glass  tubing  in  pieces  about  2  inches  long,  make  flanges  on  both  ends 
by  heating  in  a  Bunsen  flame  and  pressing  against  a  flat  piece  of 
iron  or  stone,  and  then  fasten  cheese-cloth  over  the  ends.  A  dozen 
or  more  may  be  washed  at  one  time  by  placing  them  in  a  pan  and 
allowing  water  from  the  tap  to  flow  into  the  pan.  There  should  be 
holes  in  the  bottom  of  the  pan  so  that  about  half  the  water  will  flow 
through  the  holes  rather  than  over  the  rim  of  the  pan.  Some  use 
little  bags  of  cheese-cloth  instead  of  the  glass  tubes. 

If  running  water  is  not  available,  put  the  material  into  a  rather 
large  bottle  or  dish;  a  200  c.c.  bottle  is  not  too  large  for  half  a 
dozen  j-inch  cubes.  Change  frequently,  especially  at  first.  Nothing 
is  safe  with  less  than  24  hours  of  this  sort  of  washing. 

If  the  washing  has  not  been  thorough,  the  subsequent  staining 
is  likely  to  be  unsatisfactory. 

Some  of  the  chromic-acid  formulae  are  as  follows: 

a)  Stock  Chromo-Acetic  Solution. — 

Chromic  acid 1  g. 

Glacial  acetic  acid 1  c.c. 

Water 100  c.c. 

This  solution  has  been  used  quite  extensively  in  embryological 
work  upon  the  higher  plants.  It  fixes  thoroughly,  but  often  causes 
plasmolysis  in  cells  with  large  vacuoles. 

6)  Weak  Chromo-Acetic  Solution  (Shaffner's  formula).— 

Chromic  acid 0 . 3  g. 

Acetic  acid 0 . 7  g. 

Water 99.0  c.c. 

This  has  also  been  used  in  embryological  work.  It  causes  little 
or  no  plasmolysis.  Difficult  material,  like  Aster  heads  and  ripe 


Reagents  25 

Capsella  pods,  cuts  more  readily  after  this  reagent  than  after  the 
stronger  solution. 

c)  Medium  Chromo-Acetic  Solution. — 

Chromic  acid 0 . 5  g. 

Glacial  acetic  acid 1.0  c.c. 

Water 100.0  c.c. 

This  is  a  useful  formula.  The  chromic  is  too  strong  for  some 
algae,  but  for  fern  prothallia  and  most  liverworts  the  solution  is 
quite  successful. 

d)  Flemming's  Fluid  (stronger  solution) : 

f  1  per  cent  chromic  acid 45  c.c. 

\  Glacial  acetic  acid 3  c.c. 

B.     2  per  cent  osmic  acid 12  c.c. 

Keep  the  mixture  A  made  up,  and  add  B  as  the  reagent  is  needed 
for  use,  since  it  does  not  keep  well.  This  fluid  is  quite  expensive 
on. account  of  the  osmic  acid.  For  cytological  work  it  has  been  very 
popular,  and  it  is  especially  recommended  for  chromosomes,  centro- 
somes,  achromatic  structures,  and  mitotic  phenomena  in  general. 
The  fluid  should  be  allowed  to  act  for  24  to  48  hours  and  the  washing 
in  water  must  be  very  thorough. 

Material  should  be  in  very  small  pieces  £  inch  square,  or  in  thin 
slices  £  inch  or  less  in  thickness,  for  the  fluid  penetrates  poorly.  The 
blackening  due  to  the  osmic  acid  may  be  removed  by  peroxide  of 
hydrogen  just  before  the  slide  is  passed  from  the  alcohol  into  the 
stain.  Harper  and  Holden,  in  their  work  on  Coleosporium,  recom- 
mended 4  hours  on  the  slide  in  a  3  per  cent  solution  of  the  peroxide 
of  hydrogen.  Some  prefer  a  stronger  solution  of  the  peroxide  of 
hydrogen,  even  20  per  cent.  The  peroxide  should  be  in  water,  if  one  is 
following  it  by  an  aqueous  stain,  but  may  be  in  50  per  cent  alcohol 
if  it  is  to  be  followed  by  an  alcoholic  stain.  Yamanouchi  has  used 
chlorine  for  bleaching,  and  the  results  are  fully  equal  to  those  obtained 
with  peroxide  of  hydrogen,  and  the  chlorine  is  cheaper.  Make 
the  bleacher  as  follows:  Place  some  potassium  chlorate  crystals — a 
group  about  as  large  as  a  grain  of  wheat — in  the  bottom  of  a  100  c.c. 
Stender  dish;  add  one  drop  of  25  per  cent  hydrochloric  acid  in  water; 


26  Methods  in  Plant  Histology 

immediately  fill  the  Stender  full  of  30  per  cent  alcohol  and  thus 
dissolve  the  fumes  in  alcohol.  This  will  bleach  sections  in  10  minutes, 
or  even  less.  Wash  in  30  per  cent  alcohol  2  or  3  hours  before  stain- 
ing. Trondle  uses  1  per  cent  chromic  acid  in  water  for  bleaching; 
it  is  slow,  requiring  about  8  hours,  but  he  maintains  that  material 
stains  better  than  after  bleaching  with  peroxide  of  hydrogen. 
According  to  Miss  Merriman,  the  linin  in  the  nuclei  of  onion  root- 
tips  is  not  so  well  preserved  in  this  solution,  but  the  arrangement  of 
the  chromatin  granules  is  brought  out  with  greater  distinctness. 
Flemming's  safranin,  gentian-violet,  orange  combination  gives  excel- 
lent results  after  this  reagent. 

d)  Flemming's  Fluid  (weaker  solution). — 

C  1  per  cent  chromic  acid 25  c.c. 

A  j  1  per  cent  acetic  acid 10  c.c. 

I  Water 55  c.c. 

B.  1  per  cent  osmic  acid 10  c.c. 

As  in  case  of  the  stronger  solution,  mix  A  and  B  only  as  needed  for 
immediate  use. 

Many  prefer  the  weaker  solution,  because  the  blackening  is  not 
so  extreme  and  material  does  not  become  quite  so  brittle.  Some 
allow  the  solution  to  act  for  an  hour  and  then  transfer  the  material  to 
solution  A  for  about  24  hours.  This  secures  the  rapid  killing, 
which  is  the  principal  virtue  of  the  osmic  acid,  and  avoids  the  dis- 
agreeable blackening,  so  that  little  or  no  bleaching  may  be  necessary. 

e)  Benda's  Fluid.— 

1  per  cent  chromic  acid 16  c.c. 

2  per  cent  osmic  acid 4  c.c. 

Glacial  acetic  acid 2  drops 

This  modification  of  Flemming's  stronger  solution  has  been  used 
very  successfully  in  recent  investigations  upon  chromatin. 

/)  Merkel's  Fluid.— 

Equal  volumes  of  a  1.4  per  cent  solution  of  chromic  acid  and  a 
1.4  per  cent  solution  of  platinic  chloride.  This  is  also  an  expensive 
reagent.  It  is  recommended  for  mitotic  phenomena,  but  does  not 
seem  to  equal  Flemming's  solution. 


Reagents  27 

gr)  Hermann's  Fluid. — 

1  per  cent  platinic  chloride 15  parts 

Glacial  acetic  acid 1  part 

2  per  cent  osmic  acid 4  or  2  parts 

This  is  the  most  expensive  fixing  agent  yet  discovered,  and  for 
botanical  purposes  it  does  not  seem  to  be  any  better  than  the  cheaper 
chromic  mixtures.  It  is  mentioned  here  with  chromic  mixtures 
because  it  originated  as  a  variation  of  Flemming's  fluid,  the  platinic 
chloride  being  substituted  for  the  chromic  acid. 

PICRIC  ACID 

Use  a  saturated  solution  in  water  or  70  per  cent  alcohol.  One 
gram  of  picric  acid  crystals  will  saturate  about  75  c.c.  of  water  or 
alcohol.  This  reagent  penetrates  well  and  does  not  make  the  material 
brittle.  It  is  to  be  recommended  when  difficulty  is  anticipated  in 
the  cutting.  If  used  cold,  the  time  varies  from  1  to  24  hours,  de- 
pending upon  the  character  of  the  tissue  and  size  of  the  specimen. 
If  "used  hot  (85°  C.),  5  or  10  minutes  will  be  sufficient.  Material 
should  be  washed  in  70  or  50  per  cent  alcohol.  Water  is  injurious, 
and  some  even  go  so  far  as  to  avoid  aqueous  stains,  unless  the  ma- 
terial has  been  thoroughly  washed.  The  washing  should  be  continued 
until  the  material  appears  whitish  and  the  alcohol  no  longer  becomes 
tinged  with  yellow.  Picro-carmine  gives  its  best  result  after  this  re- 
agent. Picric  acid  can  be  combined  with  various  other  fixing  agents, 
and  so  we  have  picro-sulphuric  acid,  picro-nitric  acid,  picro-chromic 
acid,  picro-chromic-sulphuric  acid,  picro-osmic  acid,  picro-alcohol, 
and  picro-corrosive  sublimate.  The  picric  acid  in  all  mixtures 
should  be  rather  strong. 

A  picric-acid  combination  which  has  gained  some  popularity 
for  cytological  work  is 

Bouin's  Fluid.— 

Formalin  (commercial) 25  c.c. 

Picric  acid  (saturated  solution  in  water) 75  c.c. 

Glacial  acetic  acid 5  c.c. 

Fix  about  24  hours.  Rinse  in  water  for  a  few  minutes  to  remove 
the  more  superficial  picric  acid,  and  then  complete  the  washing  in 


28  Methods  in  Plant  Histology 

35  per  cent  or  50  per  cent  alcohol.  There  is  likely  to  be  some 
swelling,  but  spindles  of  mitotic  figures  stain  well.  It  would  be 
worth  while  to  try  this  solution  with  embryo-sacs  of  angiosperms 
and  with  early  stages  in  the  female  gametophyte  of  gymnosperms. 

CORROSIVE  SUBLIMATE 

Corrosive  sublimate,  or  bichloride  of  mercury,  is  soluble  in  water 
and  in  alcohol.  About  5  g.  will  make  a  saturated  solution  in  100  c.c. 
of  water.  It  is  somewhat  more  soluble  in  alcohol,  but  for  practical 
purposes  5  g.  in  100  c.c.  of  50  per  cent  alcohol  may  be  regarded  as  a 
saturated  solution.  Corrosive  sublimate  used  alone  does  not  give 
as  good  results  as  when  mixed  with  acetic  acid,  chloroform,  or  picric 
acid.  Fixing  is  very  rapid,  the  material  being  fixed  almost  as  soon 
as  it  is  penetrated  by  the  fluid.  Material  which  is  at  all  transparent, 
like  some  ovules  and  the  endosperm  of  gymnosperms  before  the 
formation  of  starch,  becomes  opaque  as  soon  as  fixed,  and  so  the  time 
needed  for  fixing  is  easily  determined.  From  10  minutes  to  one 
hour  should  be  sufficient  for  onion  root-tips  or  lily  ovaries.  Smaller 
or  larger  objects  require  shorter  or  longer  periods.  When  used  hot 
(85°  C.),  the  fixing  is  much  more  rapid.  Filamentous  algae  or  fungi 
are  simply  dipped  into  the  fixing  agent  and  immediately  taken  out. 
One  minute  is  enough  for  onion  root-tips,  and  two  minutes  is  enough 
for  a  lily  ovary  at  the  fertilization  period. 

Wash  out  aqueous  solutions  with  water  and  alcoholic  solutions 
with  alcohol.  In  either  case,  the  washing  must  be  very  thorough, 
since  preparations  from  incompletely  washed  material  are  sure  to  be 
disfigured  by  crystals  of  corrosive  sublimate.  After  material  fixed 
in  the  alcoholic  solution  has  been  washed  in  alcohol  for  several  hours, 
add  to  the  50  per  cent  alcohol  a  little  of  the  iodine  solution  used 
in  testing  for  starch.  It  will  impart  a  brownish  color  to  the  alcohol, 
but  the  color  will  disappear  in  a  few  seconds,  and  the  alcohol  will 
become  clear  if  any  corrosive  sublimate  remains.  Add  more  and 
more  iodine  until  the  brown  color  fails  to  disappear.  The  washing  is 
then  complete. 

Material  fixed  in  aqueous  solutions  should  be  passed  through  the 
alcohols — as  described  under  "Dehydrating  Agents,"  a  few  pages 
farther  on — before  using  the  iodine. 


Reagents  29 

Camphor  may  be  used  instead  of  iodine  to  hasten  the  washing,  but 
it  does  not  give  any  color  reaction. 

Material  should  be  imbedded  as  soon  as  possible,  since  it  gets 
brittle  if  allowed  to  remain  in  alcohol. 

Kinoplasmic  structures  do  not  stain  well  with  gentian-violet, 
but  safranin  and  the  haematoxylins  stain  almost  as  well  as  after 
chromic-acid  mixtures,  and  the  carmines  give  their  most  brilliant 
stains,  as  a  result  of  the  formation  of  mercuric  carminate. 

The  following  formulae  are  merely  suggestive : 

a)  Corrosive  Sublimate  and  Acetic  Acid. — 

Corrosive  sublimate 3  g. 

Glacial  acetic  acid 3  c.c. 

Alcohol  (or  water) 100  c.c. 

6)  Corrosive  Sublimate,  Acetic  Acid,  and  Picric  Acid. — 

Corrosive  sublimate 5  g. 

Glacial  acetic  acid 5  c.c. 

Picric  acid,  saturated  solution  in  50  per  cent 
alcohol 100  c.c. 

c)  Corrosive  Sublimate  and  Picric  Acid  (Jeffrey's  solution). — 
Corrosive  sublimate,  saturated  solution  in  30 

per  cent  alcohol 3  parts 

Picric  acid,  saturated  solution  in  30  per  cent 

alcohol 1  part 

It  would  be  worth  while  to  try  other  combinations. 

FORMALIN 

Formalin  is  an  excellent  preservative.  It  has  been  mentioned 
already  as  an  ingredient  in  several  formulae.  Commercial  formalin 
has  a  strength  of  40  per  cent.  Throughout  this  book,  a  2,  4,  or  6  per 
cent  formalin  is  understood  to  mean  2,  4,  or  6  c.c.  of  commercial 
formalin  to  98,  96,  or  94  c.c.  of  water,  alcohol,  or  any  other  ingredient. 
Commercial  formalin  is  sure  to  contain  some  formic  acid.  For  most 
purposes,  it  is  neither  necessary  nor  desirable  to  remove  the  acid. 
For  studying  the  origin  of  vacuoles,  it  is  necessary  to  have  neutral 
formalin,  which  can  be  secured  from  commercial  formalin  by  dis- 
tillation. Place  some  sodium  bicarbonate  in  a  flask  and  distil  by 


30  Methods  in  Plant  Histology 

heating  over  a  Bunsen  flame.  It  is  not  worth  while  to  distil  more 
than  is  needed  for  immediate  use,  since  the  formic  acid  soon  reappears. 

For  filamentous  algae  and  fungi  a  3  to  6  per  cent  solution  of  the 
ordinary  commercial  formalin  in  water  is  very  good.  Material  is 
left  in  the  solution  until  needed  for  use.  For  marine  algae  sea-water 
should  be  used  instead  of  fresh  water.  Both  marine  and  fresh- 
water material  should  be  washed  for  half  an  hour  in  fresh  water  before 
staining.  A  6  per  cent  solution  will  fix  one-fourth  its  volume  of 
material.  With  material  like  filamentous  algae  or  leafy  liverworts, 
a  10  per  cent  solution  will  fix  all  one  can  put  into  the  bottle  without 
crowding. 

For  class  use,  material  should  be  washed  in  water  for  several 
minutes,  because  the  fumes  are  irritating  to  the  eyes  and  mucous 
membranes. 

For  a  study  of  the  origin  of  vacuoles  the  following  combination 
is  recommended: 

Bensley's  Formula. — 

1.  Formalin  (neutral) 10 .0  c.c. 

2.  Bichromate  of  potash 2 . 5  g. 

3.  Corrosive  sublimate 5 .0  g. 

4.  Water 90.0  c.c. 

Make  the  solution  2,  3,  4,  and  then  add  the  neutral  formalin. 
Fix  about  24  hours.  Wash  in  water,  but  use  the  iodine — necessary 
on  account  of  the  corrosive  sublimate — just  before  staining  sections 
on  the  slide. 

GENERAL  HINTS  ON  FIXING 

It  is  very  desirable  that  the  fixing  agent  penetrate  quickly  to  all 
parts  of  the  object.  For  this  reason  material  should  be  in  small 
pieces.  The  best  fixing  agents  do  their  best  work  near  the  surface 
of  the  piece.  Of  course,  filamentous  algae  and  fungi,  and  delicate 
objects  like  fern  prothallia  and  root-tips,  are  simply  thrown  into  the 
fixing  agent.  Alcohol,  formalin  alcohol,  or  formalin  alone,  may 
penetrate  |-inch  cubes;  but  the  chromic-acid  series,  which  gives 
the  best  results  in  cytological  work,  penetrates  so  poorly  that  cells 
more  than  -fa  inch  from  the  surface  are  not  likely  to  be  well  fixed. 
Most  objects  should  be  trimmed  with  a  razor  so  that  no  part  shall 


Reagents  31 

be  more  than  y1^  inch  from  the  surface.  Even  then,  it  must  be  remem- 
bered that  a  waxy  or  cutinized  or  suberized  surface  presents  an  almost 
impassable  barrier  to  the  chromic  series. 

Some  objects,  although  small,  cause  trouble  in  various  ways. 
Many  buds  are  hairy  and  will  not  sink;  if  such  things  are  dipped 
quickly  in  strong  alcohol,  they  will  usually  sink.  If  rather  large 
air  bubbles  prevent  the  material  from  sinking,  as  in  case  of  peri- 
chaetical  leaves  of  some  mosses  and  involucral  leaves  of  liverworts,  a 
little  dissection  or  a  careful  snip  with  the  scissors  will  obviate  the 
difficulty.  If  an  air-pump  is  available  some  bubbles  are  easily 
removed,  but  air  bubbles  in  cells  may  resist  even  the  air-pump. 
Heating  followed  by  rapid  cooling  is  recommended  by  Pfeiffer  and 
Wellheim  for  removing  air,  but,  for  cytological  work,  the  remedy  is 
worse  than  the  bubbles. 

It  is  often  asked  whether  fixing  agents  really  preserve  the  actual 
structure  of  cell  contents.  It  must  be  admitted  that  some  things — 
notably  the  liquid  albuminoids — are  much  modified  in  appearance, 
but  the  most  competent  observers  are  now  inclined  to  believe  that 
such  delicate  objects  as  chromosomes,  centrosomes,  the  achromatic 
figure,  and  even  the  structure  of  protoplasm,  can  be  studied  with 
confidence  from  material  which  has  been  fixed,  imbedded,  and 
stained.  Extensive  investigations  upon  various  objects  in  the  living 
condition  have  strengthened  this  confidence. 

It  is  certain  that  we  have  not  yet  found  the  ideal  fixing  agent 
for  cell  contents.  Such  an  agent  must  not  be  a  solvent  of  any  of  the 
cell  contents,  must  penetrate  rapidly,  must  preserve  structures 
perfectly,  and  must  harden  so  thoroughly  that  every  detail  shall 
remain  unchanged  during  the  subsequent  processes  of  dehydrating, 
clearing,  imbedding,  sectioning,  and  staining. 

DEHYDRATING  AGENTS 

Objects  which  are  to  be  imbedded  in  paraffin  or  celloidin,  and 
also  all  other  objects  which  are  to  be  mounted  in  balsam,  must  be 
dehydrated,  i.e.,  they  must  be  freed  from  water.  The  slightest 
trace  of  water  is  ruinous.  Alcohol  is  used  almost  exclusively  for 
dehydrating.  The  process  must  be  gradual.  If  material  has  been 


32  Methods  in  Plant  Histology 

fixed  in  an  aqueous  solution,  it  must  pass  through  a  series  of  alcohols 
of  increasing  strength,  beginning  with  about  3  per  cent  alcohol.  Ten 
years  ago,  most  botanists  were  beginning  with  35  per  cent  alcohol; 
in  the  second  edition  of  this  book  (1905)  we  recommended  15,  35, 
50,  70,  85,  95,  and  100  per  cent  as  a  safe  series,  since  it  causes  no 
obvious  plasmolysis  of  the  cell  contents.  As  investigations  have 
become  more  and  more  critical,  especially  investigations  upon  the 
structure  of  chromatin,  it  has  been  found  that  even  15  per  cent 
alcohol  is  too  strong  for  a  beginning.  It  is  maintained  that,  in 
addition  to  the  damage  done  by  transferring  from  water  to  so  strong 
an  alcohol,  the  final  dehydration  is  not  so  perfect  as  it  is  when  the 
series  begins  with  a  weaker  alcohol.  Yamanouchi,  whose  work 
upon  delicate  algae  has  been  particularly  successful,  uses  the  following 
series:  2£,  5,  7£,  10,  15,  20,  30,  40,  50,  70,  85,  95,  and  100  per  cent. 
After  such  gradual  early  stages,  there  seems  to  be  no  objection  to  the 
less  gradual  stages  which  follow.  Of  course,  there  is  no  particular 
virtue  in  the  fractions:  it  is  convenient  to  make  a  10  per  cent  alcohol, 
then  dilute  it  one-half  for  the  5  per  cent,  and  dilute  the  5  per  cent  one- 
half  for  the  2|  per  cent.  The  7|  per  cent  is  made  with  sufficient 
accuracy  by  adding  a  little  water  to  the  10  per  cent  alcohol.  For 
each  of  the  first  four  or  five  grades,  3  or  4  hours  is  long  enough.  It 
is  a  good  plan  to  change  morning,  noon,  and  night.  Fr"om  the  20  to 
the  95,  change  morning  and  evening.  About  24  hours,  changing 
two  or  three  times,  is  not  too  long  for  the  absolute  alcohol.  The 
grades  of  alcohol  below  100  per  cent  can  be  used  several  times.  The 
absolute  alcohol  should  not  be  used  again  for  this  purpose,  but  it 
should  be  saved  and  used  for  rinsing  slides  after  the  paraffin  ribbons 
have  been  dissolved  off  with  xylol  or  turpentine.  Even  85  and  95 
per  cent  alcohol  will  be  useful  for  rinsing  one's  hands  when  dealing 
with  Venetian  turpentine.  If  it  is  necessary  to  be  very  economical, 
the  stronger  alcohols  may  be  filtered  into  a  single  large  bottle  and 
the  strength  of  the  mixture  can  then  be  determined  by  using  an 
alcoholometer.  Knowing  the  strength  of  the  mixture,  one  can 
easily  make  any  weaker  grade. 

Be  sure  that  the  bottles  or  Stenders  for  absolute  alcohol  are 
perfectly  dry;  keep  the  bottles  well  corked  and  keep  the  lids  on 


Reagents 


the  Stenders.  The  importance  of  excluding  moisture  cannot  be 
exaggerated. 

The  lower  grades  are  made  up  from  95  per  cent  alcohol. 

Formulae  for  Alcohols. — The  following  formulae  will  enable 
anyone  to  make  the  other  grades  of  alcohol  from  95  per  cent  alcohol 
and  water. 

95  95  95  95  95  95  95  95  95 
101520304050607085 
85  80  75  65  55  45  35  25  10 

The  foregoing  are  the  formulae  for  various  alcohols  from  10  to 
85  per  cent.  The  first  column  shows  the  formula  for  making  10  per 
cent  alcohol.  The  percentage  of  alcohol  secured  in  each  case  is 
indicated  by  the  middle  number  in  each  column.  In  the  first  formula, 
subtract  10  from  95;  the  result,  85,  is  the  number  of  cubic 
centimeters  of  water  which  must  be  added  to  10  c.c.  of 
95  per  cent  alcohol  in  order  to  obtain  10  per  cent  alcohol. 
The  mixture  contains  95  c.c.  of  10  per  cent  alcohol.  If 
more  or  less  than  95  c.c.  of  the  mixture  is  needed,  take 
proportional  parts  of  10  and  85.  This  simple  method  is 
a  time-saver,  but  if  the  bottles  or  Stender  dishes  are  to 
be  filled  frequently,  it  will  be  a  still  further  saving  of 
time  to  use  a  long  label  (Fig.  13)  and,  after  pouring  in 
the  95  per  cent  alcohol,  draw  a  line  showing  how  high  it 
reaches,  and  then,  after  pouring  in  the  water,  draw 
another  line.  The  next  time  it  is  necessary  to  fill  the 
bottles  merely  pour  in  95  per  cent  alcohol  until  it 
reaches  the  first  line,  and  then  pour  in  water  until  it 
reaches  the  second  line.  It  is  not  necessary  to  use  distilled  water  if 
pure  drinking-water  is  available. 

Synthol  is  used  like  alcohol,  and  many  believe  it  to  be  a  good 
substitute. 

Some  investigators  use  more  or  less  complicated  diffusion  appa- 
ratus and  make  the  dehydration  process  extremely  gradual.  Judging 
from  the  finished  preparation,  we  find  no  advantage  in  the  method. 
In  the  diffusion  process,  the  solution  is  constantly  changing.  This 
may  not  be  an  advantage. 


70 
per  cent 


FIG.    13.— 
Label    for 

staining-dish. 


34  Methods  in  Plant  Histology 

Some  very  minute  objects,  like  bacteria  and  the  smaller  Cyano- 
phyceae,  may  be  dehydrated  by  heating  them  until  all  water  is 
drawn  off,  but,  of  course,  this  shows  merely  the  form,  with  little  or 
nothing  of  the  internal  structure. 

CLEARING  AGENTS 

Clearing  agents  are  so  named  because  they  render  objects  trans- 
parent. When  clearing  agents  are  used  to  precede  infiltration  with 
paraffin,  the  clearing  is  merely  incidental,  the  real  purpose  being  to 
replace  the  dehydrating  agent  with  a  solvent  of  paraffin.  The 
clearing  is  useful,  even  in  this  case,  because  it  indicates  when  the 
replacing  has  become  complete. 

When  the  clearing  agent  is  used  to  precede  infiltration  with 
paraffin,  the  material  should  always  be  most  thoroughly  dehydrated 
with  absolute  alcohol  before  beginning  with  the  clearing  agent. 
When  the  clearing  agent  is  used  to  clear  sections  or  small  objects  just 
before  mounting  in  balsam,  absolutely  perfect  dehydration  is  not 
necessary  with  all  clearing  agents.  Bergamot  oil,  carbolic  acid,  and 
Eycleshymer's  clearing  fluid  (equal  parts  of  bergamot  oil,  carbolic 
acid,  and  cedar  oil)  will  clear  readily  from  95  per  cent  alcohol. 
Sections  to  be  cleared  in  xylol  or  clove  oil  should  be  dehydrated  in 
absolute  alcohol. 

Xylol. — In  our  opinion,  xylol  is  the  best  clearing  agent  to  pre- 
cede infiltration  with  paraffin.  After  the  material  has  been  dehy- 
drated, it  should  be  brought  gradually  into  xylol.  Twenty  years 
ago  it  was  customary  to  bring  material  directly  from  absolute 
alcohol  into  xylol;  ten  years  ago,  two  or  three  mixtures  of  absolute 
alcohol  and  xylol  were  used  before  reaching  the  pure  xylol;  at 
present,  those  who  are  doing  the  most  critical  work  are  making  this 
process  still  more  gradual.  As  cytologists  have  been  studying  more 
and  more  minute  structures,  the  methods  have  become  more  and 
more  critical.  As  in  the  case  of  the  alcohol  series,  the  xylol  series 
has  its  grades  closer  together  at  the  beginning  than  at  the  end. 
The  following  series  seems  to  be  sufficiently  gradual :  -^ ,  f ,  \,  f ,  f ,  pure 
xylol.  It  is  hardly  necessary  to  use  a  graduate  in  making  up  the 
series.  For  the  f ,  use  equal  parts  of  xylol  and  absolute  alcohol;  for 


Reagents  35 

the  |,  use  equal  parts  of  the  |  and  absolute  alcohol;  for  the  f,  use 
equal  parts  of  the  %  and  absolute,  and  for  the  y1^ ,  equal  parts  of  the  f 
and  absolute.  The  f  can  be  guessed  at  with  sufficient  accuracy. 
About  4  hours  in  each  grade  is  long  enough;  change  morning, 
noon,  and  night.  The  pure  xylol  should  be  changed  once  or  twice. 
While  the  pure  xylol  must  not  be  used  again  for  this  purpose,  it  is 
still  good  for  dissolving  paraffin  ribbons  when  staining  on  the 
slide. 

Xylol  is  the  best  agent  for  clearing  sections  just  before  mounting 
in  balsam.  Preparations  cleared  in  xylol  harden  more  rapidly, 
and  this  is  such  a  decided  advantage  that  even  when  sections  have 
been  cleared  in  cedar  oil  or  clove  oil  it  is  worth  while  to  give  them  a 
minute  or  two  in  xylol  before  mounting. 

Xylol  evaporates  so  rapidly  that  one  must  take  care  not  to  let 
sections  become  dry  before  applying  the  balsam.  Thin  sections 
perfectly  dehydrated  will  clear  in  a  few  seconds;  a  minute  or  two 
should  be  sufficient  for  sections  20  /x  in  thickness.  If  there  is  much 
moisture  in  the  air,  or  if  the  absolute  alcohol  is  not  above  suspicion, 
clear  sections  in  clove  oil  before  transferring  to  xylol. 

Chloroform. — Some  botanists  use  chloroform  to  precede  the 
infiltration  with  paraffin.  In  the  later  stages  of  infiltration  it  is  more 
easily  removed  than  xylol.  It  seems  to  possess  no  other  advantages, 
and  for  clearing  sections  just  before  mounting  in  balsam  it  is  inferior 
to  xylol  or  clove  oil.  Its  value  in  hardening  celloidin  and  as  a  fixing 
agent  entitles  it  to  a  place  in  the  histological  laboratory. 

Cedar  Oil. — It  is  not  always  easy  to  get  good  cedar  oil.  If  the 
stuff  offered  for  sale  looks  like  turpentine  and  smells  like  it,  it  is 
worthless  for  histological  purposes.  Good  cedar  oil  has  a  slightly 
amber  tint,  the  color  resembling  a  weak  clove  oil.  It  should  have 
the  pleasant  odor  of 'cedar  wood.  The  very  expensive  cedar  oil 
used  with  immersion  lenses  is  not  needed  for  clearing  or  for  preced- 
ing infiltration  with  paraffin.  It  is  claimed  that  material  cleared  in 
cedar  oil  does  not  become  so  brittle  as  that  cleared  in  xylol  or  chloro- 
form. 

Clove  Oil. — This  is  an  excellent  agent  for  clearing  sections  and 
small  objects  just  before  mounting  in  balsam.  It  clears  more  readily 


36  Methods  in  Plant  Histology 

than  xylol.  When  the  absolute  alcohol  has  deteriorated  so  that  xylol 
no  longer  clears  the  sections,  clove  oil  may  still  clear  with  ease. 
While  clove  oil  will  clear  from  95  per  cent  alcohol,  it  is  better  to  use 
absolute.  Since  preparations  cleared  in  clove  oil  harden  slowly,  it  is 
a  good  plan  to  treat  them  with  xylol  before  mounting  in  balsam. 
Gentian-violet  is  somewhat  soluble  in  clove  oil,  and  this  fact  makes 
it  possible  to  secure  a  beautiful  differentiation,  because  the  stain 
is  extracted  from  some  elements  more  rapidly  than  from  others. 
The  stain  may  be  extracted  completely  from  the  chromosomes 
during  the  metaphase  and  still  remain  bright  in  the  achromatic 
structures.  After  the  desired  differentiation  has  been  attained,  the 
preparation  should  be  placed  in  xylol  to  remove  the  clove  oil,  since 
the  continued  action  of  the  clove  oil  would  cause  the  preparation  to 
fade.  Do  not  use  a  Stender  dish  for  clove  oil,  but  keep  it  in  a  50  c.c. 
bottle.  Put  on  a  few  drops,  and  immediately  drain  them  off  in  such 
a  way  as  to  remove  the  alcohol  as  completely  as  possible.  Then 
flood  the  slide  and  pour  the  clove  oil  back  into  the  bottle,  repeating 
the  process  until  the  proper  differentiation  has  been  reached.  Re- 
place the  clove  oil  with  xylol  and  mount  in  balsam.  With  stains 
not  soluble  in  clove  oil,  the  xylol  is  not  necessary,  except  to  facilitate 
the  hardening  of  the  preparation. 

.  Clove  oil  is  used  in  removing  the  celloidin  matrix  from  celloidin 
sections.  It  is  useless  as  an  agent  to  precede  infiltration  with  paraffin. 

Eycleshymer's  Clearing  Fluid. — This  is  a  mixture  of  equal  parts 
of  bergamot  oil,  cedar  oil,  and  carbolic  acid.  It  clears  readily  from 
95  per  cent  alcohol,  and  consequently  is  useful  in  clearing  celloidin 
sections  when  it  is  desirable  to  preserve  the  celloidin  matrix.  In 
sections  stained  with  haematoxylin,  or  haematoxylin  and  eosin,  the 
stain  may  be  removed  completely  from  the  matrix  by  the  use  of 
acid  alcohol,  and  the  matrix  may  be  preserved  by  clearing  from  95 
per  cent  alcohol. 

It  is  not  intended  that  the  mixture  should  be  used  to  precede 
infiltration  with  paraffin. 

Other  Clearing  Agents. — Bergamot  oil,  carbolic  acid,  turpentine, 
benzine,  gasoline,  and  other  reagents  have  been  tried  for  clearing, 
but  none  seem  to  be  worth  more  than  a  warning  mention. 


Reagents  37 

MISCELLANEOUS  REAGENTS 

Canada  Balsam  is  used  almost  exclusively  for  mounting.  Very 
thick  balsam  is  disagreeable  to  handle  and  makes  unsatisfactory 
mounts.  Very  thin  balsam,  in  drying  out,  allows  bubbles  to  run 
under  the  cover.  Xylol  is  cheaper  than  balsam,  and  consequently 
the  balsam  on  the  market  is  likely  to  be  too  thin  for  immediate  use. 
The  stopper  may  be  left  out  until  the  balsam  acquires  the  proper 
consistency.  Material  cleared  in  clove  oil  or  cedar  oil  may  be 
mounted  directly  in  xylol  balsam.  It  is  not  necessary  that  the  clear- 
ing agent  should  be  also  the  solvent  of  the  balsam. 

Paraffin  should  be  of  at  least  two  grades,  a  soft  paraffin  melting 
at  40°  to  45°  C.,  and  a  hard  paraffin  melting  at  52°  to  54°  C. 
Griibler's  paraffin  and  most  imported  paraffins  melt  at  the  tempera- 
ture indicated  on  the  wrappers.  The  melting-point  indicated  on  the 
wrappers  of  paraffins  sold  by  some  American  dealers  does  not  enable 
one  to  make  even  a  guess  as  to  the  real  melting-point.  One  promi- 
nent optical  company  sells  a  paraffin  marked  70°  C.,  which  usually 
melts  at  52°  to  55°  C.  The  fact  that  the  price  rises  with  the  melting- 
point  may  explain  the  discrepancy. 

Paraffin  may  be  used  repeatedly.  Keeping  it  in  the  liquid  con- 
dition in  the  bath  month  after  month  is  an  advantage,  since  it 
becomes  more  and  more  tenacious  and  homogeneous. 

Glycerin,  glycerin  jelly,  Venetian  turpentine,  and  gold  size  are 
described  in  the  chapter  on  "The  Glycerin  Method"  (chap.  vii). 
Celloidin  is  described  in  the  chapter  on  "The  Celloidin  Method" 
(chap.  x).  The  reagents  already  described  are  noted  further  in  con- 
nection with  specific  applications.  Reagents  used  in  making  micro- 
chemical  tests  are  described  in  the  chapter  on  "  Temporary  Mounts 
and  Microchemical  Tests"  (chap.  v). 

A  list  of  reagents  with  suggestions  in  regard  to  quantities  and 
prices  will  be  found  in  chap.  xxix. 


CHAPTER  III 
STAINS  AND  STAINING 

During  the  past  ten  years  no  new  stains  of  the  first  rank  have 
come  into  favor,  but  much  greater  precision  has  been  attained  in  the 
use  of  some  which  were  already  popular.  For  cytological  work 
Haidenhain's  iron-haematoxylin  holds  a  firm  place  at  the  head  of  the 
list,  with  Flemming's  triple  stain  an  easy  second.  For  anatomical 
work,  safranin  still  holds  first  place  for  the  lignified  elements  of  the 
vascular  system,  but  the  claim  of  Delafield's  haematoxylin  to  first 
place  for  cellulose  tissues  is  no  longer  undisputed,  for  anilin  blue 
is  giving  excellent  results  and  light  green  (Licht  Grun,  as  it  reads 
on  the  label)  seems  to  give  more  accurate  views  of  the  phloem 
than  we  were  securing  with  any  of  the  other  stains.  The  fact  that 
excellent  preparations  can  be  made,  almost  without  trial,  by  using 
combinations  already  perfected  doubtless  deters  investigators  from 
experimenting  with  other  stains.  There  is  still  abundant  room  for 
experimenting  with  various  stains,  and  especially  in  the  use  of  mor- 
dants and  in  the  effect  of  the  same  stain  or  combination  after  various 
fixing  agents.  It  is  to  be  regretted  that  botanists  who  need  micro- 
technic  have  so  little  knowledge  of  chemistry,  and  that  chemists 
have  no  interest  in  developing  methods  of  staining. 

Stains  may  be  classified  in  various  ways:  e.g.,  there  are  three 
great  groups  of  stains — the  carmines,  the  haematoxylins,  and  the 
anilins.  Stains  may  be  classified  as  basic  and  acid,  or  they  may  be 
regarded  as  general  and  specific.  A  general  stain  affects  all  the  ele- 
ments, while  a  specific  stain  affects  only  certain  elements,  or  stains 
some  elements  more  deeply  than  others.  Stains  which  show  a 
vigorous  affinity  for  the  nucleus  have  been  called  nuclear  stains,  and 
those  which  affect  the  cytoplasm  more  than  the  nucleus  have  been 
termed  plasma  stains.  Of  course,  such  stains  are  specific. 

We  shall  consider  some  of  the  more  important  haematoxylins, 
carmines,  and  anilins,  reserving  general  directions  and  theoretical 

38 


Stains  and  Staining  39 

questions  for  another  chapter.  The  formulae  are  largely  empirical. 
Some  of  those  given  here  are  taken  from  The  Microtomist's  Vade- 
Mecum  (Lee),  which  is  easily  the  most  complete  compendium  of 
stains  and  other  reagents  concerned  in  microtechnic.  It  is  to  be 
regretted  that  botanists  have  no  book  of  this  character,  but  it  must 
be  confessed  that  we  have  not  the  material  for  such  an  extensive 
work.  Other  formulae  are  from  Botanical  Microtechnique  (Zimmer- 
mann)  and  from  Stirling's  Histology,  and  still  others  are  from  our 
own  laboratory.  The  directions  for  using  a  stain  apply  to  stains 
made  up  according  to  the  formulae  which  are  given  here,  and  may 
need  modification  if  other  formulae  are  employed.  It  is  hoped,  how- 
ever, that  the  directions  will  give  the  student  sufficient  insight  into 
the  rationale  of  staining  to  enable  him  to  make  any  necessary  modi- 
fications. 

The  current  practice  in  staining  paraffin  sections  on  the  slide 
differs  from  the  practice  in  staining  freehand  sections  or  small 
objects  which  are  to  be  mounted  whole.  In  case  of  paraffin  sections, 
the-  cell  contents  are  usually  as  important  and  often  more  important 
than  the  cell  walls;  consequently,  extreme  care  must  be  given  to 
every  detail.  With  freehand  sections  the  cell  contents  often  drop 
out,  but  even  when  they  remain  the  cell  walls  are  usually  the  impor- 
tant features;  and  so  the  process  is  considerably  shortened. 

For  staining  freehand  sections,  it  is  customary  to  use  solid 
watch  glasses,  unless  the  sections  are  very  large.  The  details  of 
the  method  are  given  in  chap,  vi,  on  "  Freehand  Sections." 

For  staining  sections  on  the  slide,  nothing  is  better  than  the 
ordinary  Stender  dish.  The  arrangement  of  Stender  dishes  shown 
in  Fig.  14  is  very  convenient.  The  advantage  is  obvious.  With 
two  dishes  each  of  xylol,  xylol-alcohol,  and  absolute  alcohol,  one 
set  can  be  used  in  passing  down  to  the  stain,  and  the  other,  which 
is  thus  kept  free  from  any  paraffin  in  solution,  can  be  used  in  passing 
back  to  the  balsam.  Even  for  paraffin  sections,  some  use  only 
three  alcohols,  50,  95,  and  100  per  cent,  and  the  first  two  may  be 
simply  poured  over  the  slide;  in  this  case,  only  one  Stender  dish — 
for  the  100  per  cent  alcohol — is  necessary  in  the  alcohol  series,  the 
other  two  alcohols  being  kept  in  bottles.  This  short  method  gained 


40  Methods  in  Plant  Histology 

great  popularity  because  it  was  used  in  Stras.burger's  laboratory  at 
Bonn.  It  was  the  influence  of  this  school  and  its  great  master 
which  led  to  the  adoption  of  the  short  schedule  in  the  second  edition 
of  this  book.  A  few  years'  trial  showed  the  weakness  of  the  method, 
and  we  returned  to  the  longer  schedule.  The  crudeness  of  the 
short  schedule  is  doubtless  responsible  for  the  tenacity  with  which 
the  Bonn  school  has  clung  to  the  theory  of  linin  and  chromomeres. 
The  young  investigator  should  be  warned  that  during  the  last 
twenty  years  of  his  life,  Strasburger,  who  had  been  a  leader  in 


FIG.  14. — A  convenient  arrangement  of  staining-dishes 

technic,  cut  very  few  sections  and  did  practically  no  staining,  but 
used  preparations  made  by  assistants. 

Let  us  now  consider  a  few  of  the  most  important  stains. 

THE  HAEMATOXYLINS 

The  most  important  haematoxylins  are  Haidenhain's  iron-alum 
haematoxylin,  Delafield's  haematoxylin,  Mayer's  haem-alum,  and 
Boehmer's  haematoxylin. 

All  the  haematoxylins  mentioned  contain  alum,  and,  according 
to  Mayer,  who  has  written  the  most  important  work  on  haematoxylin 
stains,1  "the  active  agent  in  them  is  a  compound  of  haematin  with 
alumina.  This  salt  is  precipitated  in  the  tissues,  chiefly  in  the  nuclei, 
by  organic  and  inorganic  salts  there  present  (e.g.,  by  the  phosphates), 
and  perhaps  also  by  other  organic  bodies  belonging  to  the  tissues." 
These  salts  are  fixed  in  the  tissues  by  the  killing  and  fixing  agent, 
and  when  the  stain  is  applied  a  chemical  combination  results. 
Haematoxylins  stain  well  after  any  of  the  fixing  agents  described  in 

«  "  Ueber  das  Farben  mit  Hamatoxylin,"  MMheilungen  aus  der  Zoologischen  Station  zu 
Neapel,  10  : 170-186,  1891,  and  "  Ueber  Hamatoxylin,  Carmin  und  verwandten  Materien," 
Zeitschri/t  fur  wissenschaftliche  Mikroscopie,  16  : 196-220,  1899. 


Stains  and  Staining  41 

the  preceding  paper,  but  they  are  most  effective  when  used  after 
members  of  the  chromic-acid  series. 

Haidenhain's  Iron- Alum  Haematoxylin. — This  stain  was  intro- 
duced by  Haidenhain  in  1892  and  has  gained  a  well-deserved  popu- 
larity with  those  engaged  in  cytological  work.  Two  solutions  are 
used,  and  they  are  never  mixed: 

A.  2  to  4  per  cent  aqueous  solution  of  ammonia  sulphate  of  iron. 

B.  \  per  cent  aqueous  solution  of  haematoxylin. 

In  making  solution  A,  use  the  violet  ferric  crystals,  not  the  ferrous. 

The  first  solution  acts  as  a  mordant,  i.e.,  it  does  not  stain,  but 
prepares  the  tissue  for  the  action  of  the  second  solution. 

Solution  A  is  at  its  best  as  soon  as  the  crystals  are  completely 
dissolved  and  it  remains  in  practically  perfect  condition  for  about 
two  months,  after  which  it  gradually  deteriorates. 

The  haematoxylin  crystals  for  solution  B  should  be  dissolved  in 
water.  This  will  require  about  10  days.  The  solution  should  then 
be  .allowed  to  "  ripen"  for  4  weeks  before  it  is  ready  for  use.  Unfor- 
tunately, it  remains  at  its  best  for  only  a  short  time,  not  more  than 
5  or  6  weeks.  This  is  because  the  "ripening,"  which  is  an  oxidation 
process,  continues,  and  the  solution  becomes  too  ripe.  Some  prefer 
to  dissolve  the  haematoxylin  crystals  in  alcohol — about  10  g.  in 
100  c.c.  of  absolute  alcohol.  This  solution  should  stand  until  it  has 
a  deep  wine-red  color.  This  will  require  4  or  5  months,  and  a  year  is 
not  too  long.  From  this  stock  solution,  make  up  small  quantities 
as  needed.  About  4  or  5  c.c.  of  this  stock  solution  in  100  c.c.  of 
water  gives  a  practically  aqueous  solution,  and  it  is  already  ripe. 

The  general  method  is  as  follows:  treat  with  A,  stain  in  B,  and 
then  return  to  A  to  reduce  and  differentiate  the  stain.  Never 
transfer  directly  from  A  to  B,  or  from  B  to  A;  always  wash  in  water 
before  passing  from  one  of  the  solutions  to  the  other.  It  is  a  good 
plan  to  use  a  4  per  cent  solution  of  A  to  precede  the  stain  and  a  2 
per  cent  solution  for  differentiating. 

While  all  follow  the  general  method  just  indicated,  no  two  investi- 
gators would  prepare  exactly  the  same  schedule,  even  for  staining  the 
same  object,  e.g.,  root-tips;  neither  investigator  would  use  the  same 
schedule  for  a  root-tip  and  an  embryo-sac;  an  alga  might  require 


42  Methods  in  Plant  Histology 

different  treatment,  and  all  the  preceding  variations  might  fail 
miserably  with  the  pollen  tubes  of  cycads.  This  stain  is  so  impor- 
tant that  every  worker  must  learn  it,  and  the  only  way  to  learn 
it  is  to  become  acquainted  with  the  general  outline  of  the  process 
and  then  adapt  every  step  to  the  case  in  hand. 

For  the  sake  of  illustration,  I  asked  two  prominent  cytologists, 
Dr.  S.  Yamanouchi  and  Dr.  L.  W.  Sharp,  both  of  whom  have  been 
notably  successful  in  staining  mitotic  figures,  to  write  schedules 
indicating  their  methods  of  using  this  stain.  While  both  protested 
that  the  practice  could  not  be  written  down,  they  kindly  prepared 
the  following  schedules,  not  for  the  instruction  of  their  colleagues,  but 
to  introduce  the  method  to  beginners.  Both  schedules  are  for 
paraffin  sections.  Throughout  the  first  schedule,  I  have  interpolated 
comments  and  suggestions. 

Yamanouchi's  Schedule.— 

1.  Xylol,  5  minutes,  to  dissolve  the  paraffin. 

Do  not  heat  the  slides  to  melt  the  paraffin.  However,  a  gentle 
warming  which  does  not  approach  the  melting-point  of  the  paraffin 
does  no  damage  and  makes  the  paraffin  dissolve  more  readily.  The 
xylol  soon  has  considerable  paraffin  in  solution,  but  100  c.c.  of  xylol 
should  remove  the  paraffin  from  at  least  100  slides  with  ribbons 
25  mm.  long  and  10  p  thick.  If  the  ribbons  are  only  5  ft  thick, 
200  slides  can  be  treated. 

2.  Xylol  and  absolute  alcohol,  equal  parts,  5  minutes. 

3.  Absolute  alcohol,  5  to  7  minutes. 

4.  95,  85,  70,  50,  35  per  cent  alcohol,  5  minutes  each. 

If  material  has  been  fixed  in  a  reagent  containing  osmic  acid, 
it  should  be  bleached.  For  this  purpose,  10  to  15  c.c.  of  hydrogen 
peroxide  may  be  added  to  100  c.c.  of  the  50  per  cent  alcohol. 

5.  Water,  10  to  20  minutes. 

If  any  alcohol  is  left  in  the  sections,  the  staining  will  not  be 
brilliant.  Change  the  water  several  times. 

6.  Iron-alum. 

Use  the  4  per  cent  solution.  For  many  objects,  like  the  arche- 
gonia  of  gymnos perms  and  the  embryo-sacs  of  angiosperms,  1  hour  is 
usually  enough.  For  chromosomes  in  root-tips  and  anthers,  2  hours 
may  be  long  enough;  but  for  algae,  2  hours  is  generally  a  minimum. 

7.  Wash  in  water,  5  minutes. 

The  water  should  be  changed  several  times.  If  the  washing  is 
not  thorough,  the  differentiation  will  not  be  sharp. 


Stains  and  Staining  43 

8.  Haematoxylin. 

Many  objects,  like  the  archegonia  of  gymnosperms  and  the 
embryo-sacs  of  angiosperms,  will  stain  sufficiently  in  5  or  6  hours; 
algae  require  at  least  20  hours. 

9.  Wash  in  water,  5  minutes,  changing  as  often  as  the  water  shows 
any  color. 

10.  Iron-alum,  2  per  cent  solution. 

No  time  can  be  indicated  here.  The  preparation  must  be 
watched  under  the  microscope.  After  some  experience,  one  can 
form  some  judgment  from  the  color  tone,  as  the  slide  stands  in  the 
Stender  dish  of  iron-alum,  but  the  finishing  must  always  be  done 
under  the  microscope.  If  the  stain  is  coming  out  rather  slowly,  as 
it  should,  one  can  handle  6  to  10  slides  at  one  time.  Put  the  slides 
on  a  5X7  glass  plate  and  put  the  plate  on  the  stage  of  the  micro- 
scope. The  iron-alum  can  be  added  or  removed  with  a  pipette. 
As  slide  after  slide  reaches  the  proper  differentiation,  it  is  placed 
in  water. 

11.  Water,  30  minutes. 

The  water  should  be  changed  several  times.  If  this  washing 
is  not  thorough,  the  preparation  will  fade,  on  account  of  the  con- 
tinued action  of  the  iron-alum.  If  an  aqueous  counter-stain  is 
used,  apply.it  at  this  point. 

12.  35,  50,  70,  85,  95,  100  per  cent  alcohol,  5  minutes  in  each. 

If  an  alcoholic  counter-stain  is  used,  apply  it  near  the  alcohol  of 
the  same  strength  as  the  stain. 

13.  Absolute  alcohol  and  xylol,  equal  parts,  5  minutes. 

14.  Xylol,  2  to  5  minutes. 

15.  Balsam. 

Sharp's  Schedule.— 

1.  Remove  the  paraffin  with  xylol. 

2.  Rinse  in  absolute  alcohol. 

3.  95  per  cent  alcohol. 

4.  50  per  cent  alcohol. 

5.  Water. 

6.  If  osmic  acid  has  been  used  in  fixing,  place  the  slides  in  10  per  cent 
solution  of  peroxide  of  hydrogen  in  water  until  bleached. 

7.  Water. 

8.  Iron-alum,  1\  per  cent,  2  to  3  hours. 

9.  Wash  well. 

10.  \  per  cent  haematoxylin,  24  hours. 

11.  Wash  in  water. 

12.  Extract  the  stain  in  1  per  cent  iron-alum,  watching  the  process 
under  the  microscope. 


44  Methods  in  Plant  Histology 

13.  Wash  several  hours  in  water. 

14.  Alcohol  series:  10,  30,  50,  70,  80,  95,  100  per  cent. 

If  a  counter-stain  is  desired,  introduce  it  in  one  of  the  alcohols 
of  this  series. 

15.  Absolute  alcohol  and  xylol,  equal  parts. 

16.  Xylol. 

17.  Mount  in  balsam. 

While  these  two  schedules  would  enable  the  student  to  apply 
the  method  in  case  of  objects  to  be  mounted  whole,  like  filamentous 
algae,  fern  prothallia,  etc.,  a  complete  schedule  is  given  in  chap. 
viii  on  "The  Venetian  Turpentine  Method." 

The  times  given  above  must  not  be  accepted  as  final.  Many 
prefer  to  wash  in  water  for  several  hours  after  the  first  immersion  in 
iron-alum.  Some  think  that  4  hours  is  enough  for  the  entire  process. 
Many  put  the  slide  into  iron-alum  in  the  morning  and  finish  the 
process  in  the  afternoon.  These  short  schedules  are  not  likely  to 
prove  satisfactory  with  mitotic  figures.  A  plan  which  has  proved 
convenient  and  very  successful  is  to  put  the  slide  into  the  iron-alum 
in  the  morning,  let  it  wash  in  water  during  the  afternoon,  leave  it  in 
the  \  per  cent  of  haematoxylin  over  night,  and  finish  the  prepara- 
tion the  next  morning.  It  is  a  long  process,  requiring  care,  patience, 
and  judgment,  but  it  is  worth  the  effort. 

Chromosomes,  centrosomes,  and  pyrenoids  take  a  brilliant  black, 
or,  if  the  second  treatment  with  iron-alum  be  more  prolonged, 
a  blue  black  or  purple.  Achromatic  structures  stain  purple,  but 
the  stain  can  be  extracted  while  it  is  still  bright  in  the  chromosomes. 
Lignified,  suberized,  and  cutinized  structures  stain  lightly  or  not  at 
all.  Cellulose  does  not  stain  so  deeply  as  with  Delafield's  haema- 
toxylin. Archesporial  cells  and  early  stages  in  sporogenous  tissue 
stain  gray.  Many  details  which  are  not  so  brilliantly  colored  often 
show  good  definition. 

If  a  counter-stain  is  desired,  anything  which  gives  a  serviceable 
contrast  may  be  used. .  In  any  case,  the  haematoxylin  stain  must  be 
complete  and  the  washing  thorough  before  the  second  stain  is  applied. 
An  aqueous  stain  should  be  applied  just  after  the  final  washing  in 
water;  an  alcoholic  stain  should  be  applied  during  the  process  of 
passing  the  slides  through  the  alcohols,  staining  in  a  solution  of  saf- 
ranin  in  50  per  cent  alcohol  from  the  35  or  50  per  cent  alcohol;  and 


Stains  and  Staining  45 

staining  after  the  final  absolute  alcohol,  if  the  stain  is  dissolved  in 
clove  oil. 

A  stain  of  3  or  4  minutes  in  safranin  adds  an  excellent  differentia- 
tion in  case  of  many  algae  and  does  not  obscure  nuclear  details. 
The  exine  of  pollen  grains  may  take  a  brilliant  red  with  safranin 
in  5  to  10  minutes,  contrasting  sharply  with  the  mouse  gray  of  the 
intine.  Orange  G,  in  clove  oil,  often  gives  a  pleasing  contrast. 

Delafield's  Haematoxylin.— "To  100  c.c.  of  a  saturated  solution 
of  ammonia  alum  add,  drop  by  drop,  a  solution  of  1  g.  of  haema- 
toxylin  dissolved  in  6  c.c.  of  absolute  alcohol.  Expose  to  air  and 
light  for  one  week.  Filter.  Add  25  c.c.  of  glycerin  and  25  c.c.  of 
methyl  alcohol.  Allow  to  stand  until  the  color  is  sufficiently  dark. 
Filter,  and  keep  in  a  tightly  stoppered  bottle"  (Stirling  and  Lee). 
The  addition  of  the  glycerin  and  methyl  alcohol  will  precipitate 
some  of  the  ammonia  alum  in  the  form  of  small  crystals.  The  last 
filtering  should  take  place  4  or  5  hours  after  the  addition  of  the 
glycerin  and  methyl  alcohol. 

-The  solution  should  stand  for  at  least  two  months  before  it  is 
ready  for  using.  This  "ripening"  is  brought  about  by  the  oxida- 
tion of  haematoxylin  into  haematin,  a  reaction  which  may  be  secured 
in  a  few  minutes  by  a  judicious  application  of  peroxide  of  hydrogen. 
However,  we  prefer  to  let  the  haematoxylin  ripen  naturally.  There 
is  no  objection  to  making  this  stain  in  considerable  quantity,  since 
it  does  not  deteriorate.  We  have  used  Delafield's  haematoxylin 
which  had  been  in  a  cork-stoppered  bottle  for  twenty  years,  and 
it  still  gave  the  rich  characteristic  stain. 

Transfer  to  the  stain  from  50  or  35  per  cent  alcohol  or  from  water. 
The  length  of  time  required  is  exceedingly  variable.  Sometimes 
sections  will  stain  deeply  in  3  minutes,  but  it  is  often  necessary  to 
stain  for  30  minutes  or  even  longer.  This  stain  may  be  diluted 
with  several  times  its  own  volume  of  water;  when  this  is  done,  the 
time  required  is  correspondingly  long,  but  the  staining  is  frequently 
more  precise.  The  length  of  time  required  will  be  fairly  uniform  for 
all  material  taken  from  the  same  bottle.  This  fact  indicates  that  the 
washing  process,  which  follows  killing  and  fixing,  is  an  important 
factor;  if  the  washing  has  been  thorough,  the  material  will  stain 
readily;  but  if  the  washing  has  been  insufficient,  the  material  may 


46  Methods  in  Plant  Histology 

stain  slowly  or  not  at  all.  The  washing  is  particularly  important 
when  the  fixing  agent  contains  an  acid.  Transfer  from  the  stain 
to  tap  water.  Distilled  water  is  neither  necessary  nor  desirable. 
Some  writers  recommend  washing  for  24  hours,  but  this  is  entirely 
unnecessary;  for  paraffin  sections  on  the  slide,  5  or  10  minutes  is 
long  enough,  and  even  for  rather  thick  freehand  sections  20  to  30 
minutes  is  sufficient.  Use  plenty  of  water  and  keep  changing  it  as 
often  as  it  becomes  in  the  least  discolored.  Precipitates  are  often 
formed  when  slides  are  transferred  directly  to  alcohol  from  this 
stain,  and  sometimes  even  after  washing  in  water.  A  few  gentle 
dips  in  acid  alcohol  (2  drops  of  HC1  to  100  c.c.  of  70  per  cent  alcohol) 
will  usually  remove  the  precipitates.  This  extracts  the  stain  more 
rapidly  from  other  parts  than  from  the  nuclei,  and  hence  gives  a  good 
nuclear  stain,  while  at  the  same  time  it  removes  any  disfiguring 
precipitates.  Some  prefer  to  stain  for  a  very  short  time  and  use  no 
acid  alcohol,  but,  as  a  rule,  it  is  better  to  overstain  and  then  differen- 
tiate in  this  way,  because  sharper  contrasts  are  obtained.  Transfer 
from  acid  alcohol  to  70  per  cent  alcohol  and  leave  here  until  a  rich 
purple  color  replaces  the  red  due  to  the  acid.  Since  small  quantities 
of  the  acid  alcohol  are  carried  over  into  the  70  per  cent  alcohol,  it 
is  well  to  add  a  drop  of  ammonia  now  and  then  to  neutralize  the 
effect  of  the  acid.  Too  much  ammonia  is  to  be  avoided,  for  it  gives 
a  disagreeable  bluish  color  with  poor  differentiation,  probably  on 
account  of  the  precipitation  of  alumina.  The  preparation  is  now 
dehydrated  in  95  per  cent  and  then  in  absolute  alcohol,  cleared  in 
xylol  or  clove  oil,  and  mounted  in  balsam. 

The  following  is  a  general  schedule  for  staining  paraffin  sections 
on  the  slide  in  Delafield's  haenlatoxylin : 

1.  Stain  (from  water  or  from  35  or  50  per  cent  alcohol) . .  10  minutes 

2.  Rinse  in  water 10  minutes 

3.  35  and  50  per  cent  alcohol 5  minutes  each 

4.  Acid  alcohol 5  seconds 

5.  70  per  cent  alcohol 5  minutes 

6.  85  per  cent  alcohol 5  minutes 

7.  95  per  cent  and  100  per  cent  alcohol 5  minutes  each 

8.  Xylol  and  100  per  cent  alcohol,  equal  parts 5  minutes 

9.  Xylol. 

10.  Mount  in  balsam. 


Stains  and  Staining  47 

If,  after  rinsing  in  water,  the  stain  is  evidently  too  weak,  put 
the  slide  or  section  back  into  the  stain  until  it  appears  overstained. 
Place  the  slide  in  acid  alcohol.  If  an  acid  alcohol  with  2  drops  of 
HC1  to  100  c.c.  of  70  per  cent  alcohol  reduces  the  stain  too  much  in  4 
or  5  seconds,  use  less  acid.  Transfer  to  70  per  cent  alcohol  without 
any  acid.  As  soon  as  the  color  changes  from  red  to  purple,  examine 
under  the  microscope.  If  it  is  still  overstained,  return  to  the  acid 
alcohol;  if  the  stain  is  too  weak,  return  to  the  haematoxylin  and 
try  it  again.  After  the  haematoxylin  is  just  right,  apply  a  contrast 
stain,  if  you  wish  to  double  stain.  Before  transferring  to  the  xylol 
wipe  the  alcohol  from  the  back  of  the  slide,  or  at  least  rest  the  corner 
of  the  slide  upon  blotting-paper  for  two  or  three  seconds,  in  order 
that  you  may  not  carry  over  so  much  alcohol  into  the  xylol.  Add 
a  drop  of  balsam  and  a  cover.  Since  the  xylol  is  very  volatile,  this 
last  step  must  be  taken  quickly.  If  blackish  spots  appear  they 
are  usually  caused  by  the  drying  of  sections  before  the  balsam  and 
cover  are  added;  if  there  are  whitish  spots  or  an  emulsion-like 
appearance,  the  clearing  is  not  thorough;  this  may  be  caused  by 
poor  xylol  (or  other  clearing  agent);  by  absolute  alcohol  which  is 
considerably  weaker  than  its  name  implies  (the  absolute  alcohol 
must  test  at  least  as  high  as  99  per  cent,  and  ought  to  test  as  high 
as  99.5  per  cent,  if  xylol  is  to  be  used  for  clearing);  or  by  passing 
too  quickly  through  the  absolute  alcohol  and  xylol,  or  even  by 
moisture  on  the  cover-glass.  The  last  danger  is  easily  avoided  by 
passing  the  cover  quickly  through  a  Bunsen  or  alcohol  flame  before 
laying  it  on  the  balsam. 

Delafield's  haematoxylin  is  the  most  generally  useful  stain  in  the 
haematoxylin  group.  It  brings  out  cellulose  walls  very  sharply, 
and  consequently  is  a  good  stain  for  embryos  and  the  fundamental 
tissue  system  in  general.  With  safranin  it  forms  a  good  combination 
for  the  vascular  system,  the  safranin  giving  the  lignified  elements  a 
bright  red  color,  while  the  haematoxylin  stains  the  cellulose  a  rich 
purple.  It  is  a  good  stain  for  chromatin,  and  the  achromatic  struc- 
tures show  up  fairly  well,  but  can  be  brought  out  much  better  by 
special  methods.  Archesporial  cells  and  sporogenous  tissue  are 
very  well  defined  if  proper  care  be  taken.  Lignified  and  suberized 


48  Methods  in  Plant  Histology 

walls  and  also  starch  and  chromatophores  stain  lightly  or  not  at  all. 
Whenever  you  are  in  doubt  as  to  the  selection  of  a  stain  for  general 
purposes,  we  should  advise  the  use  of  Delafield's  haematoxylin, 

Mayer's  Haem-Alum. — Haematoxylin,  1  g.,  dissolved  with  gentle 
heat  in  50  c.c.  of  9*5  per  cent  alcohol  and  added  to  a  solution  of  50  g. 
of  alum  in  a  liter  of  distilled  water.  Allow  the  mixture  to  cool  and 
settle;  filter;  add  a  crystal  of  thymol  to  preserve  from  mold  (Lee). 

It  is  ready  for  use  as  soon  as  made  up.  Unless  attacked  by  mold, 
it  keeps  indefinitely.  Transfer  to  the  stain  from  water.  It  is  seldom 
necessary  to  stain  for  more  than  10  minutes,  and  4  or  5  minutes  is 
generally  long  enough.  As  a  rule,  better  results  are  secured  by 
diluting  the  stain  (about  1  c.c.  to  10  c.c.  of  distilled  water)  and 
allowing  it  to  act  for  10  hours  or  over  night. 

This  is  a  good  stain  for  the  nuclei  of  filamentous  algae  and  fungi, 
since  it  has  little  or  no  effect  upon  cell  walls  or  plastids.  Wash 
thoroughly  in  water  and  transfer  to  10  per  cent  glycerin.  Specimens 
may  be  mounted  in  Venetian  turpentine,  as  described  in  chap.  viii. 

Erlich's  Haematoxylin. — 

Distilled  water 50  c.c. 

Absolute  alcohol 50  c.c. 

Glycerin 50  c.c. 

Glacial  acetic  acid 5  c.c. 

Haematoxylin 1  g. 

Alum  in  excess. 

Keep  it  in  a  dark  place  until  the  color  becomes  a  deep  red.  If 
well  stoppered,  it  will  keep  indefinitely.  Transfer  to  the  stain  from 
50  per  cent  or  35  per  cent  alcohol.  Stain  5  to  30  minutes.  Since 
there  is  no  danger  from  precipitates  and  the  solution  does  not  over- 
stain,  it  is  not  necessary  to  treat  with  water  or  with  acid  alcohol,  but 
the  slide  may  be  transferred  from  the  stain  to  70  per  cent  alcohol. 
Eosin,  erythrosin,  or  orange  G  are  good  contrast  stains.  Jeffrey 
uses  safranin  and  Erlich's  haematoxylin  for  woody  tissues. 

Boehmer's  Haematoxylin. — 

f  Haematoxylin 1  g. 

\  Absolute  alcohol 12  c.c. 

•n  f  Alum 1  g. 

I  Distilled  water. .  .  .240  c.c. 


Stains  and  Staining  49 

The  solution  A  must  ripen  for  two  months.  When  wanted  for 
use,  add  about  10  drops  of  A  to  10  c.c.  of  B.  Stain  10  to  20  minutes. 
Wash  in  water  and  proceed  as  usual. 

Cellulose  walls  take  a  deep  violet.  The  closing  membrane 
(torus)  of  the  bordered  pits  of  conifers  will  usually  stain  deeply  in 
about  15  minutes.  Lignified,  suberized,  and  cutinized  structures 
stain  slightly  or  not  at  all.  When  they  do  stain,  the  color  is  not 
violet,  but  a  light  yellow  or  brown. 

THE  CARMINES 

This  group  of  stains,  immensely  popular  several  years  ago,  has 
rapidly  lost  favor  among  botanists  as  newer  stains  and  combinations 
have  appeared.  Botanists  have  not  given  the  carmines  a  fair  trial 
in  recent  years.  It  is  possible  that  it  would  be  worth  while  to  try 
again,  especially  after  fixing  agents  containing  mercury.  When  it  is 
desirable  to  stain  in  bulk,  nothing  has  been  found  which  will  serve 
better  than  the  carmines.  Only  three  of  these  stains  will  be  con- 
sidered : 

Greenacher's  Borax  Carmine. — 

Carmine 3  g. 

Borax 4  g. 

Distilled  water 100  c.c. 

Dissolve  the  borax  in  water  and  add  the  carmine,  which  is 
quickly  dissolved  with  the  aid  of  gentle  heat.  Add  100  c.c.  of  70 
per  cent  alcohol  and  filter  (Stirling). 

The  following  is  a  slightly  different  method  for  making  this  stain 
from  the  ingredients  mentioned  above:  Dissolve  the  borax  in  water, 
add  the  carmine,  and  heat  gently  for  10  minutes;  after  the  solution 
cools,  add  the  alcohol  and  filter;  let  the  solution  stand  for  2  or  3 
weeks,  then  decant  and  filter  again. 

Stain  the  material  in  bulk  from  50  per  cent  alcohol  1  to  3  days, 
then  treat  with  acid  alcohol  (50  c.c.  of  70  per  cent  alcohol +2  drops 
of  hydrochloric  acid)  until  the  color  becomes  a  clear  red;  this  may 
require  only  a  few  hours,  but  may  take  2  or  3  days.  The  material 
may  then  be  passed  through  the  rest  of  the  alcohols  (6  to  24  hours 
each),  cleared,  imbedded,  and  cut.  After  the  sections  are  fastened 


50  Methods  in  Plant  Histology 

to  the  slide,  the  paraffin  should  be  dissolved  off  with  xylol.  The 
balsam  and  cover  may  be  added  immediately,  or  the  xylol  may  be 
rinsed  off  with  alcohol  and  a  contrast  stain  may  be  added. 

Alum  Carmine. — A  4  per  cent  aqueous  solution  of  ammonia  alum 
is  boiled  20  minutes  with  1  per  cent  of  powdered  carmine.  Filter 
after  it  cools  (Lee). 

Stain  from  12  to  24  hours  and  wash  in  water.  No  acid  alcohol 
is  needed,  since  the  solution  does  not  overstain. 

Alum  Cochineal. — 

Powdered  cochineal 50  g. 

Alum 5  g. 

Distilled  water 500  c.c. 

Dissolve  the  alum  in  water,  .add  the  cochineal,  and  boil;  evapo- 
rate down  to  two-thirds  of  the  original  volume,  and  filter.  Add  a 
few  drops  of  carbolic  acid  to  prevent  mold  (Stirling). 

Stain  as  with  alum  carmine.  It  used  to  be  a  common  practice 
to  stain  in  bulk  in  alum  cochineal  and  counter-stain  on  the  slide 
with  Bismarck  brown. 

THE  ANILINS 

Many  of  the  most  brilliant  and  beautiful  stains  yet  discovered 
belong  to  this  group.  These  stains  are  so  numerous  that  we  shall 
not  attempt  to  mention  even  their  names,  but  shall  consider  only 
those  which  are  in  most  common  use  by  botanists.  The  following 
general  formula  has  proved  to  be  fairly  satisfactory  for  most  anilins, 
but  the  formulae  mentioned  in  describing  the  different  stains  are 
usually  to  be  preferred.  Solutions  containing  anilin  oil  do  not  keep 
as  well  as  aqueous  or  alcoholic  solutions. 

General  Formula. — Make  a  10  per  cent  solution  of  anilin  oil  in 
95  per  cent  alcohol;  when  the  anilin  oil  is  dissolved,  add  enough  water 
to  make  the  whole  mixture  about  20  per  cent  alcohol;  add  1  g.  of 
cyanin,  erythrosin,  safranin,  gentian-violet,  etc.,  to  each  100  c.c.  of 
this  solution. 

The  anilins  keep  well  in  balsam  but  not  in  glycerin.  Xylol 
is  a  good  clearing  agent  for  all  of  them,  but  clove  oil  should  be  used 
with  gentian-violet.  Unfortunately  they  are  not  very  permanent. 


Stains  and  Staining  51 

Preparations  fade  rapidly  if  exposed  to  bright  sunlight.  Keep  the 
slides  in  the  box  when  not  in  use,  and,  even  when  in  use,  do  not  leave 
them  scattered  over  the  laboratory  tables,  exposed  to  bright  light. 

Some  of  the  anilins  are  acid,  some  basic,  and  some  are  neutral. 

The  rapidity  with  which  sections  must  be  transferred  from  one 
fluid  to  another  makes  many  of  them  more  difficult  to  manage  than 
the  haematoxylins  or  the  carmines,  but  the  stains  are  so  valuable 
that  even  the  beginner  should  spend  most  of  his  time  with  the  anilins. 

Many  anilins  stain  quite  deeply  in  1  to  20  minutes,  but  if  the 
stain  washes  out  during  the  dehydrating  process,  stain  longer,  even 
10  to  24  hours  if  necessary.  Often  the  brilliancy  of  the  stain  can  be 
increased  by  leaving  the  slide  for  5  minutes  in  a  1  per  cent  solution  of 
permanganate  of  potassium  before  staining.  The  permanganate 
acts  as  a  mordant. 

The  following  are  the  more  important  anilins  now  in  use  by 
botanists.  The  directions  apply  to  solutions  made  up  according  to 
the -formulae  given  with  the  different  stains. 

Safranin. — Two  safranins  are  sold  by  dealers,  one  soluble  in 
water  and  the  other  soluble  in  alcohol.  The  alcoholic  is  somewhat 
soluble  in  water  and  the  aqueous  is  somewhat  soluble  in  alcohol, 
but  both  make  better  solutions  when  used  with  their  intended 
solvents. 

The  best  aqueous  solution  is  simply  a  1  per  cent  solution  in 
distilled  water. 

The  alcoholic  solution  is  made  by  dissolving  1  g.  of  the  alcoholic 
safranin  in  100  c.c.  of  95  per  cent  or  absolute  alcohol  and,  after  the 
safranin  is  completely  dissolved,  adding  50  c.c.  of  distilled  water. 

According  to  Flemming,  dissolve  0.5  g.  of  alcoholic  safranin  in 
50  c.c.  of  absolute  alcohol,  and  after  4  days  add  10  c.c.  of  distilled 
water. 

A  method  which  we  have  used  for  more  than  ten  years  with  good 
results  is  to  make  a  1  per  cent  solution  of  the  aqueous  safranin  in 
distilled  water;  then  make  a  1  per  cent  solution  of  the  alcoholic 
safranin  in  95  per  cent  alcohol;  then  mix  equal  volumes  of  the  two 
solutions.  This  makes  a  strong  solution  of  safranin  in  about  50  per 
cent  alcohol. 


52  Methods  in  Plant  Histology 

An  anilin  safranin  may  be  made  according  to  the  general 
formula. 

The  transfer  to  the  stain  depends  upon  the  formula.  If  the  stain 
is  aqueous,  transfer  to  the  stain  from  water;  if  made  up  according  to 
the  general  anilin  oil  formula,  transfer  to  the  stain  from  water  or, 
if  coming  down  from  higher  alcohols,  from  35  per  cent  alcohol;  if 
the  mixture  of  aqueous  and  alcoholic  safranins  is  used,  transfer  from 
35  per  cent  alcohol,  if  going  up  in  the  series,  and  from  70  per  cent 
alcohol,  if  coming  down  from  stronger  alcohols.  For  freehand 
sections  of  woody  tissues  we  always  use  the  mixture.  If  sections  are 
cut  from  living  material,  leave  them  in  95  per  cent  alcohol  for  half 
an  hour  and  then  transfer  to  the  stain.  Sections  cut  from  alcoholic 
material  may  be  transferred  directly  to  the  stain.  If  cut  from 
formalin-alcohol  material,  leave  the  sections  in  50  per  cent  or  70 
per  cent  alcohol  for  ten  minutes  before  transferring.  If  cut  from 
formalin  material,  leave  them  in  water  for  10  minutes,  then  in  35 
per  cent  alcohol  for  10  minutes  before  staining. 

The  time  required  for  staining  varies  with  the  tissue,  the  fixing 
agent,  and  the  quality  of  the  stain.  In  general,  it  may  be  said  that 
2  hours  is  a  minimum  and  24  hours  a  maximum.  If  the  staining  be 
too  prolonged,  delicate  structures,  like  starch  grains,  crystals,  and 
various  cell  constituents,  may  wash  out.  The  mere  fact  that  the 
whole  section  does  not  wash  off  does  not  mean  that  everything  is 
fastened  to  the  slide.  On  the  other  hand,  it  is  difficult  to  get  a 
sharp  differentiation.  In  staining  a  vascular  bundle,  one  should  be 
able  to  wash  the  safranin  from  the  cellulose  walls  and  still  leave  a 
brilliant  red  in  lignified  structures.  For  paraffin  sections,  3  to  6  hours 
will  usually  be  sufficient.  It  is  a  good  practice  to  put  the  slides  into 
the  stain  in  the  morning  and  finish  the  mounts  any  time  in  the  after- 
noon. For  freehand  sections  of  woody  tissues,  24  hours  is  not  too 
long. 

From  the  stain  transfer  to  50  per  cent  alcohol.  If  the  sections 
are  deeply  stained,  and  sufficient  differentiation  is  not  secured  within 
5  or  10  minutes,  a  drop  of  hydrochloric  acid  added  to  50  c.c.  of  the 
alcohol  will  hasten  the  extraction  of  the  stain.  If  staining  vascular 
tissue,  draw  the  stain  from  the  cellulose  walls,  but  stop  before  the 


Stains  and  Staining  53 

lignified  walls  begin  to  fade.  If  staining  mitotic  figures,  draw 
the  stain  from  the  spindle,  but  stop  before  the  chromosomes  begin 
to  weaken.  When  the  desired  differentiation  has  been  reached,  wash 
out  the  acid  in  50  per  cent  alcohol,  if  acid  has  been  used.  About  5 
minutes  should  be  sufficient. 

If  safranin  is  to  be  used  alone,  pass  through  50,  70,  85,  95,  and 
100  per  cent  alcohol,  through  the  xylol-alcohol,  then  through  xylol 
to  balsam.  If  clove  oil  is  used,  omit  the  xylol-alcohol,  but  follow 
the  clove  oil  with  xylol  to  hasten  the  hardening  of  the  preparation. 

If  a  second  stain  is  to  be  added,  transfer  from  the  50  per  cent 
alcohol  to  any  alcohol  stain.  If  the  second  stain  is  an  aqueous 
stain,  rinse  the  slide  or  sections  for  a  minute  in  water  before  applying 
the  stain. 

Safranin  is  the  most  generally  useful  of  all  the  red  stains,  and, 
fortunately,  it  is  quite  durable.  Lignified,  suberized,  cutinized,  and 
chitinized  structures  stain  red,  as  do  also  the  chromosomes,  nucleoli, 
and  centrosomes. 

Acid  Fuchsin. — Use  a  1  per  cent  solution  in  water  or  in  70  per  cent 
alcohol.  The  solution  in  alcohol  is  preferable  if  sections  are  to  be 
mounted  in  balsam.  This  stain  often  acts  with  great  rapidity, 
2  or  3  minutes  being  sufficient.  The  method  for  using  acid 
fuchsin  with  woody  tissues  is  given  in  the  chapter  on  "Freehand 
Sections"  (chap.  vi).  In  staining  embryo-sacs,  pollen  grains,  and 
such  structures,  longer  periods  are  better.  Stain  1  or  2  hours, 
and  then  differentiate  in  a  saturated  solution  of  picric  acid  in  70 
per  cent  alcohol.  This  may  require  30  seconds,  or  even  several 
minutes.  Rinse  in  70  per  cent  alcohol  until  a  bright  red  replaces  the 
yellowish  color  due  to  the  acid,  and  then  proceed  as  usual. 

Congo  Red.— This  is  an  acid  stain  resembling  acid  fuchsin. 
For  cytological  work  use  a  \  pjer  cent  aquetms  solution;  for 
anatomical  work  use  a  saturated  solution.  It  is  a  good  stain  to  use 
after  malachite  green  or  anilin  blue.  Transfer  to  the  Congo  red 
from  water,  stain  15  minutes,  wash  in  water,  transfer — for  wood 
sections — to  85  per  cent  alcohol,  and  wash  until  the  green  or  blue  color 
of  the  previous  stain  begins  to  show  through  the  red.  Then  treat 
quickly  with  absolute  alcohol,  clear  in  xylol,  and  mount  in  balsam. 


54  Methods  in  Plant  Histology 

Eosin. — This  has  long  been  a  favorite  stain,  but  for  most  pur- 
poses it  has  been  replaced  by  similar  stains  giving  better  differentia- 
tion. The  dry  stain  is  made  in  two  forms,  one  for  aqueous  and  the 
other  for  alcoholic  solution.  Each  should  be  used  with  its  intended 
solvent.  Make  a  1  per  cent  solution  in  alcohol  or  water. 

For  algae  or  fungi  to  be  mounted  in  glycerin  use  the  aqueous 
solution  and  stain  for  several  hours;  treat  with  1  per  cent  acetic  acid 
for  several  seconds,  and  then  wash  the  acid  out  thoroughly  in  water. 
Place  in  10  per  cent  glycerin,  and  allow  the  glycerin  to  concentrate. 
According  to  Lee,  the  glycerin  should  be  slightly  alkaline.  The 
alkalinity  can  be  brought  about  by  adding  half  a  gram  of  common  salt 
to  100  c.c.  of  the  10  per  cent  glycerin.  We  have  found  that  eosin 
keeps  well  when  the  glycerin  is  acidulated  with  about  1  c.c.  glacial 
acetic  acid  to  100  c.c.  of  10  per  cent  glycerin. 

For  staining  paraffin  sections,  the.  alcoholic  solution  is  better. 
One  or  two  minutes  is  usually  sufficient,  and  it  is  not  necessary  to 
use  acid. 

Haematoxylin  and  eosin  and  methyl  blue  and  eosin  are  good 
combinations.  The  eosin  should  follow  the  other  stain. 

Erythrosin. — This  is  really  an  eosin,  but  there  is  some  difference 
in  the  method  of  manufacturing.  It  is  a  more  precise  and  a  more 
transparent  stain  than  eosin  and  is  to  be  preferred  for  nearly  all 
staining  of  paraffin  sections.  Make  a  1  per  cent  solution  in  distilled 
water  or  in  70  per  cent  alcohol.  It  gives  good  results  when  made  up 
according  to  the  general  formula. 

Erythrosin  stains  rapidly,  30  seconds  to  3  minutes  being  sufficient. 
When  used  in  combination  with  other  stains,  erythrosin  should  come 
last. 

Magdala  Red. — At  least  two  Magdala  reds  are  sold  by  dealers, 
one  the  edit  (genuine)  Magdala  red,  and  the  other  simply  Magdala 
red.  The  latter  is  much  cheaper  and,  in  our  experience,  much 
superior  to  the  echt  stain.  The  directions  apply  to  the  cheaper  stain. 

For  staining  algae  which  are  to  be  mounted  in  Venetian  tur- 
pentine, use  a  1  per  cent  solution  in  85  or  95  per  cent  alcohol.  Dilute 
the  stain  about  one-half  with  95  per  cent  alcohol  and  allow  it  to  act 
for  6  to  8,  or  even  24,  hours.  Rinse  in  95  and  100  per  cent  alcohol 


Stains  and  Staining  55 

for  a  few  minutes.  Transfer  to  10  per  cent  Venetian  turpentine  and 
allow  the  turpentine  to  concentrate  as  described  in  chap.  viii. 

In  staining  sections  to  be  mounted  in  balsam,  the  same  stain  may 
be  used,  but  it  is  better  to  dilute  it  one-half  with  water.  Stain  for 
6  to  24  hours,  dehydrate  in  95  per  cent  and  absolute  alcohol,  clear 
in  clove  oil,  and  mount  in  balsam. 

Magdala  red  stains  lignified,  suberized,  and  cutinized  structures, 
and  also  chromosomes,  centrosomes,  nucleoli,  and  pyrenoids.  It  is 
likely  to  overstain,  but  the  differentiation  is  easily  secured  by  placing 
the  finished  mounts  upon  a  white  background  in  the  direct  sunlight. 
When  the  desired  differentiation  has  been  reached,  it  is  better  to 
avoid  direct  sunlight,  although  the  mounts  do  not  seem  to  fade  in 
the  ordinary  light  of  a  room. 

Except  for  special  purposes,  it  is  better  to  use  this  stain  in 
combination  with  blue,  green,  or  violet. 

Gentian- Violet. — This  is  one  of  the  most  important  stains  in  the 
bota'nical  laboratory.  It  may  be  made  according  to  the  general 
formula  for  anilin  stains,  but  that  solution  does  not  keep  well.  A 
1  per  cent  solution  in  distilled  water  keeps  indefinitely  and  seems  to 
be  as  good  as,  if  not  better  than,  the  anilin  solution.  Gentian- violet 
dissolves  readily  in  clove  oil,  and  this  may  prove  to  be  a  better  method 
of  making  the  stain  than  either  of  the  two  well-known  formulae. 

With  the  aqueous  or  anilin-oil  solutions,  the  following  directions 
will  enable  the  student  to  become  acquainted  with  the  behavior  of 
the  stain.  Transfer  to  the  stain  from  water  and  allow  the  stain  to 
act  for  1  to  30  minutes.  The  time  depends  upon  the  fixing  and  upon 
the  structures  to  be  stained.  The  brilliancy  of  the  stain  in  achromatic 
structures  may  often  be  increased  by  leaving  the  slide  from  2  to  5 
minutes  in  a  1  per  cent  aqueous  solution  of  permanganate  of  potas- 
sium before  applying  the  stain.  The  greatest  objection  to  the 
aqueous  and  anilin-oil  solutions  of  gentian-violet  is  that  the  stain 
washes  out  so  rapidly  in  alcohols  that  it  is  impossible  to  run  the 
slide  up  through  the  series.  The  usual  practice  is  to  dip  the  slide 
in  water  to  remove  most  of  the  stain  and  thus  avoid  carrying  it  into 
the  alcohol:  then  transfer  directly  from  water  to  95  per  cent  alcohol, 
allowing  the  alcohol  to  act  for  only  2  or  3  seconds,  then  allow  the 


56  Methods  in  Plant  Histology 

absolute  alcohol  to  act  for  5  or  6  seconds,  and  then,  while  the  stain 
is  still  coming  out  in  streams,  begin  the  treatment  with  clove  oil. 
Holding  the  slide  in  one  hand,  pour  on  a  few  drops  of  clove  oil, 
and  immediately  drain  off  in  such  a  way  as  to  carry  off  the  alcohol. 
This  clove  oil  should  not  be  used  again.  Then  flood  the  slide 
repeatedly  with  clove  oil,  pouring  the  clove  oil  back  into  the  bottle. 
A  50  c.c.  bottle  of  clove  oil  is  large  enough.  About  100  mounts  can 
be  cleared  with  50  c.c.  of  this  oil.  The  clove  oil  is  a  solvent  of 
gentian-violet,  but  it  dissolves  the  stain  from  some  structures  more 
rapidly  than  from  others;  e.g.,  the  stain  may  be  completely  removed 
from  the  chromosomes  while  it  is  still  bright  in  the  achromatic 
structures.  As  soon  as  the  stain  is  just  right,  drain  off  the  clove  oil 
and  leave  the  slide  in  xylol  for  a  minute  or  two  before  mounting  in 
balsam.  This  is  a  necessary  step,  because  the  continued  action  of 
clove  oil  would  cause  the  preparation  to  fade.  As  may  be  inferred 
from  what  has  preceded,  alcohol  would  soon  extract  the  stain,  without 
any  application  of  clove  oil.  The  clove  oil  is  used,  not  only  because 
it  extracts  the  stain  more  slowly,  but  because  it  dissolves  the  stain 
from  some  structures  more  rapidly  than  from  others;  e.g.,  the  stain 
may  be  completely  removed  from  the  chromosomes  while  it  is  still 
bright  in  the  achromatic  structures,  so  that  with  safranin  and  gentian- 
violet  one  can  get  red  chromosomes  on  a  violet  spindle. 

Some  still  use  cedar  oil  to  follow  the  clove  oil.  This  stops  the 
action  of  the  clove  oil,  but  the  preparations  harden  slowly. 

Gentian-violet  is  an  excellent  stain  for  achromatic  structures  in 
all  stages  of  development.  Chromatin,  in  many  of  its  stages,  is  also 
stained.  In  metaphase  and  anaphase  one  should  be  able  to  get 
red  chromosomes  and  violet  spindles  with  safranin  and  gentian- 
violet.  If  the  chromosomes  also  persist  in  retaining  the  violet, 
shorten  the  stain  in  gentian-violet.  Cilia  stain  well;  starch  grains 
stain  deeply,  chromatophores  less  deeply,  and  lignified  walls  may 
not  stain  at  all.  One  should  be  able  to  get  red  lignified  walls  and 
violet  cellulose  walls  with  safranin  and  gentian-violet. 

Cyanin. — This  stain  is  also  called  Quinolein  Blue  and  Chinoiin 
Blue.  Dissolve  1  g.  of  cyanin  in  100  c.c.  of  95  per  cent  alcohol  and 
add  100  c.c.  of  water.  The  cyanin  would  not  dissolve  in  50  per  cent 


Stains  and  Staining  57 

alcohol.  We  have  not  found  Griibler's  cyanin  very  satisfactory  with 
the  foregoing  formula.  With  the  general  formula  the  Griibler's  cyanin 
will  not  dissolve.  We  use  a  cyanin  prepared  by  H.  A.  Metz  &  Co., 
122  Hudson  Street,  New  York.  This  cyanin  dissolves  completely 
when  made  up  according  to  the  general  formula.  It  stains  rapidly, 
5  to  10  minutes  usually  being  sufficient.  Chromosomes  take  a  deep 
blue,  but  the  spindle  is  only  slightly  affected.  Lignified  structures 
stain  blue,  while  cellulose  walls  are  scarcely  affected  and  the  stain  is 
easily  washed  out. 

Iodine  Green. — Use  a  1  per  cent  solution  in  70  per  cent  alcohol. 
Stain  for  an  hour,  rinse  in  70  per  cent  alcohol,  dehydrate  in  95  per 
cent  alcohol  and  absolute  alcohol,  clear  in  xylol  or  clove  oil,  and 
mount  in  balsam.  If  the  stain  washes  out  too  rapidly  and  does  not 
give  sufficient  differentiation,  stain  longer,  over  night  or  even  24 
hours. 

Lignified  structures  stain  green,  but,  after  proper  washing, 
cellulose  is  scarcely  affected.  A  bright  green  may  be  left  in  the 
chromosomes  after  all  the  stain  has  been  washed  out  from  the  spindle. 

Acid  fuchsin,  erythrosin,  and  eosin  are  good  contrast  stains 
for  mitotic  figures.  Acid  fuchsin  or  Delafield's  haematoxylin  are 
good  for  cellulose  walls. 

Light  Green  (Licht  Gruri). — Light  green  is  an  acid  stain,  soluble 
in  water,  alcohol,  or  clove  oil.  It  stains  quickly  and  forms  a  sharp 
contrast  with  safranin  or  Magdala  red. 

Stain  in  safranin  and  then,  with  little  or  no  washing  out,  stain  in 
a  weak  alcoholic  solution  of  acid  green  (about  0.2  g.  in  100  c.c.  of 
95  per  cent  alcohol).  From  20  seconds  to  about  1  minute  may  be 
sufficient.  The  green  rapidly  reduces  the  safranin,  and  consequently 
the  staining  must  not  be  too  prolonged.  A  successful  preparation 
should  show  red  chromosomes  and  green  spindle.  Lignified  walls 
should  be  red  and  cellulose  walls  green. 

Malachite  Green. — A  1  to  3  per  cent  aqueous  solution  is  good  for 
cellulose  walls.  The  stain  contrasts  well  with  Congo  red. 

Methyl  Green. — A  1  per  cent  solution  in  water  is  good  for 
staining  lignified  structures.  Lee  recommends  that  the  solution 
be  acidulated  with  acetic  acid.  This  is  not  necessary  for  staining 


58  Methods  in  Plant  Histology 

lignified  membranes  nor  for  staining  chromosomes.  Methyl  green 
has  long  been  a  favorite  stain  for  living  tissues.  It  is  more  easily 
controlled  than  iodine  green,  especially  in  double  staining  to  differ- 
entiate lignified  and  cellulose  walls. 

Acid  Green. — Make  a  solution  according  to  the  general  formula, 
or  simply  make  a  1  per  cent  solution  in  water.  This  stains  cellulose 
walls  and  achromatic  structures,  but  scarcely  affects  lignified  walls  or 
chromosomes. 

Anilin  Blue. — Strong  alcoholic  solutions  are  best  for  botanical 
work.  Even  though  the  dry  stain  may  be  intended  for  aqueous 
solution,  make  a  1  per  cent  solution  in  85  or  95  per  cent  alcohol. 

This  stain  can  be  recommended  for  cellulose  walls,  achromatic 
structures  of  mitotic  figures,  for  cilia,  and  it  is  particularly  valuable 
for  algae.  Directions  for  using  it  with  algae  are  given  in  chap.  viii. 

Orange  G. — Make  a  1  per  cent  solution  in  water,  in  95  per  cent 
alcohol,  or  in  clove  oil.  We  prefer  the  solution  in  clove  oil. 

Transfer  to  the  aqueous  stain  from  water;  to  the  alcoholic  stain 
from  85  per  cent  alcohol,  since  the  stain  is  always  applied  as  a  second 
or  third  stain;  use  the  solution  in  clove  oil  after  the  dehydration  in 
absolute  alcohol.  Times  are  always  short  and  are  to  be  reckoned  in 
seconds  rather  than  in  minutes.  If  the  solution  in  clove  oil  has 
been  used,  rinse  it  off  with  pure  clove  oil  and  then  transfer  to  xylol 
before  mounting  in  balsam. 

This  is  a  plasma  stain.  It  is  distinctly  a  general  rather  than  a 
selective  stain,  but  is  valuable  as  a  background  for  other  structures 
which  have  been  stained  violet  or  blue  or  green.  If  first  came  into 
prominence  as  the  third  member  of  the  triple  stain,  safranin,  gentian- 
violet,  orange. 

Gold  Orange. — This  stain,  which  many  incorrectly  suppose  to 
be  the  same  as  orange  G,  is  much  more  readily  soluble  in  clove  oil 
and  stains  with  much  greater  rapidity. 

Bismarck  Brown. — Use  a  2  per  cent  solution  in  70  per  cent 
alcohol. 

This  is  a  good  stain  for  cellulose  walls,  although  it  is  not  so  precise 
as  haematoxylin.  Embryo-sacs  stained  in  one  of  the  carmines  are 
improved  by  1  or  2  minutes'  staining  in  Bismarck  brown.  Material 
fixed  in  alcohol  stains  better  than  that  which  has  been  fixed  in  reagents 


Stains  and  Staining  59 

containing  chromic  acid.     A  faint  background  of  Bismarck  brown  is 
quite  effective  in  staining  sections  containing  bacteria. 

Nigrosin. — Make  a  1  or  2  per  cent  solution  in  water.  A  few 
drops  of  this  solution  to  a  watch  glass  full  of  water  stains  filamentous 
algae  or  fungi  in  1  to  3  hours.  It  keeps  well  in  glycerin  or  Venetian 
turpentine.  It  also  keeps  well  in  balsam,  but  it  is  of  little  value  in 
staining  microtome  sections. 

COMBINATION  STAINS 

Sometimes  preparations  are  stained  with  a  single  stain,  selected 
to  emphasize  some  particular  feature,  but  in  the  great  majority  of 
cases  two  or  more  stains  are  used.  In  staining  a  vascular  bundle, 
one  stain  may  be  selected  which  stains  the  xylem,  but  not  the  phloem, 
while  another  of  a  different  color  stains  the  phloem,  but  not  the 
xylem,  thus  affording  a  sharp  contrast.  In  staining  mitotic  figures, 
one  stain  may  stain  the  chromosomes,  while  another  of  a  different 
color  may  be  used  to  stain  the  spindle. 

Success  in  double  staining  can  be  obtained  only  by  noting  the 
effect  of  each  stain  upon  the  various  plant  structures. 

Flemming's  Safranin,  Gentian-Violet,  Orange. — Safranin  has 
long  been  a  famous  stain  for  mitosis.  This  triple  combination  was 
published  in  1891,  but  its  value  in  plant  cytology  was  not  thoroughly 
appreciated  until  five  or  six  years  later,  when  its  application  was 
developed  to  a  high  degree  of  perfection  by  various  investigators  of 
the  Bonn  (Germany)  school.  Three  methods,  which  may  be  desig- 
nated as  A,  B,  and  C,  will  be  described. 

A.  According  to  Flemming,  stain  2  or  3  days  in  safranin  (dissolve 
0.5  g.  safranin  in  50  c.c.  absolute  alcohol,  and  after  4  days  add  10  c.c. 
distilled  water) ;  rinse  quickly  in  water;  stain  1  to  3  hours  in  a  2  per 
cent  aqueous  solution  of  gentian-violet;  wash  quickly  in  water,  and 
then  stain  1  to  3  minutes  in  a  1  per  cent  aqueous  solution  of  orange  G. 
Transfer  from  the  stain  to  absolute  alcohol,  clear  in  clove  oil,  and 
mount  in  balsam. 

B.  The  following  formulae  and  method  seem  to  be  better  for 
mitotic  phenomena  in  plants:  Make  a  1  per  cent  solution  of  alcoholic 
safranin  in  absolute  or  95  per  cent  alcohol,  and  after  the  safranin  is 
completely  dissolved,  add  an  equal  volume  of  a  1  per  cent  solution  of 


60  Methods  in  Plant  Histology 

aqueous  safranin  in  water,  thus  making  a  1  per  cent  solution  of 
safranin  in  50  per  cent  alcohol.  Use  a  1  per  cent  aqueous  solution  of 
gentian-violet,  and  a  1  per  cent  aqueous  solution  of  orange  G. 

Transfer  paraffin  sections  to  the  stain  from  95  per  cent  alcohol 
after  the  turpentine  or  xylol  used  in  dissolving  away  the  paraffin  has 
been  rinsed  off.  Stain  3  to  24  hours.  If  the  period  be  too  short, 
the  washing  out  is  so  rapid  that  it  is  difficult  to  stop  the  differentiation 
at  the  proper  point  and,  besides,  the  red  is  likely  to  be  less  brilliant. 
Rinse  in  50  per  cent  alcohol  until  the  stain  is  properly  differentiated. 
Leave  the  slide  in  the  50  per  cent  alcohol  until  the  stain  is  washed 
out  from  the  spindle  and  cytoplasm,  but  stop  the  washing  out 
before  the  chromosomes  begin  to  lose  their  bright  red  color.  If  the 
washing  out  takes  place  too  slowly,  treat  with  slightly  acidulated 
alcohol  (1  drop  of  HC1  to  50  c.c.  of  50  per  cent  alcohol)  for  a  few 
seconds.  The  acid  must  be  removed  by  washing  for  15  to  30  seconds 
in  alcohol  which  has  not  been  acidulated. 

Then  dip  the  slide  5  or  6  times  into  water  and  stain  in  gentian- 
violet.  The  time  required  is  so  variable  that  definite  instructions 
are  impossible.  The  gentian-violet  should  stain  the  spindle,  but 
not  the  chromosomes.  If  the  stain  be  too  prolonged,  it  may  be 
impossible  to  get  it  out  from  the  chromosomes  and  still  leave  it  bright 
in  the  spindle.  If  the  period  be  too  short,  the  stain  will  wash  out 
from  the  spindle.  For  mitotic  figures  in  the  germinating  spores  of  the 
liverwort,  Pellia,  30  minutes  is  not  too  long.  In  this  case,  the  stain 
washes  out  easily  from  the  chromosomes  without  the  use  of  acid,  and 
the  spindle  takes  a  rich  violet  which  is  not  easily  washed  out.  In 
embryo-sacs  of  Lilium  try  10  minutes.  In  pollen  mother  cells  try 
5  to  10  minutes.  For  root-tips  try  2  to  10  minutes.  Chromatin  in 
the  early  prophases  and  in  telophases  will  stain  with  the  violet,  and 
the  violet  will  not  wash  out,  but  in  phases  in  which  fully  formed 
chromosomes  are  visible  the  violet  can  be  washed  out  if  the  period  has 
not  been  too  long. 

Remove  the  slide  from  the  gentian-violet  and  dip  it  5  or  6  times 
in  water  and  then  stain  30  seconds  to  1  minute  in  orange  G.  The 
orange  stains  cytoplasm  and  at  the  same  time  washes  out  gentian- 
violet. 


Stains  and  Staining  61 

Transfer  from  the  orange  G  to  95  per  cent  alcohol,  dipping  the 
slide  a  few  times  in  this  merely  to  save  the  absolute  alcohol.  Dehy- 
drate in  absolute  alcohol  3  to  30  seconds. 

Clear  in  clove  oil,  as  already  described  in  the  paragraph  on 
gentian-violet.  Transfer  to  xylol  and  mount  in  balsam. 

Safranin  and  gentian-violet  are  often  used  without  the  orange. 
In  this  case,  transfer  from  the  gentian-violet  directly  to  95  per  cent 
alcohol,  and  proceed  as  before. 

A  serious  objection  to  both  these  methods  is  that  the  gradual  series 
of  alcohols  cannot  be  used,  because  the  gentian-violet  washes  out  so 
rapidly.  While  the  objection  may  be  overcome,  to  some  extent, 
by  using  the  orange  in  95  per  cent  alcohol,  solutions  of  gentian-violet 
in  strong  alcohol  have  not  been  satisfactory.  We  have  been  trying 
a  third  method,  with  more  or  less  success.  So  far,  it  seems  better 
than  either  of  the  two  methods  just  described. 

C.  Use  the  safranin  solution  described  in  B,  but  use  the  gentian- 
violet  and  orange  G  in  1  per  cent  clove-oil  solutions. 

'  For  paraffin  sections,  transfer  to  safranin  from  70  or  50  per  cent 
alcohol  and  stain  as  directed  under  B.  Pass  through  70,  85,  95, 
and  100  per  cent  alcohol,  about  5  minutes  in  each.  Put  the  clove- 
oil  gentian  solution  on  the  slide  with  a  pipette  and  stain  5  to  30 
minutes.  Drain  off  the  stain  (which  can  be  used  repeatedly),  and 
add  the  clove-oil  orange  solution  and  allow  it  to  act  10  to  20  seconds. 
This  stains  with  orange  and,  at  the  same  time,  extracts  the  gentian- 
violet.  Pour  off  the  clove-oil  orange  solution  (which,  unlike  the 
gentian  solution,  is  not  worth  much  for  a  second  staining)  and 
pour  on  pure  clove  oil.  Watch  it  until  the  gentian-violet  is  satis- 
factory, then  transfer  to  xylol  and  mount  in  balsam. 

This  method  avoids  the  big  skips  of  A  and  B,  and  the  prepara- 
tions seem  better. 

Cyanin  and  Erythrosin. — Make  both  solutions  according  to  the 
general  formula  for  anilins,  or  make  1  per  cent  aqueous  solutions, 
but  note  what  was  said  about  cyanin  in  the  paragraph  on  p.  57. 

Stain  in  cyanin  5  to  10  minutes  or  longer;  rinse  quickly  in  water 
if  using  the  aqueous  solution,  or  in  35  per  cent  alcohol  if  using  the 
general  formula;  then  stain  30  seconds  to  1  minute  in  erythrosin. 


62  Methods  in  Plant  Histology 

If  the  cyanin  washes  out,  stain  for  1  hour,  and  if  it  still  washes  out, 
omit  the  rinsing  in  alcohol  and  transfer  directly  from  the  cyanin  to 
the  eiythrosin. 

The  erythrosin  may  be  used  first;  in  this  case  stain  for  5  minutes 
in  erythrosin,  transfer  directly  to  cyanin,  and  stain  for  about  10 
seconds.  Dehydrate  in  95  per  cent  and  in  absolute  alcohol,  clear 
in  xylol  or  in  clove  oil,  and  mount  in  balsam. 

The  stains  wash  out  so  rapidly  that  the  series  of  alcohols  cannot 
be  used. 

Chromosomes  and  nucleoli  stain  blue  and  achromatic  structures 
red.  Lignified  structures  stain  blue  and  cellulose  walls  red.  The 
various  cell  constituents  are  often  sharply  differentiated.  It  was  this 
combination  which  suggested  the  now  obsolete  terms,  "cyanophi- 
lous"  and  "erythrophilous." 

Magdala  Red  and  Anilin  Blue. — Make  both  solutions  as  directed 
in  chap,  viii  on  "The  Venetian  Turpentine  Method." 

Stain  3  to  24  hours  in  Magdala  red,  dip  in  95  per  cent  alcohol  to 
rinse  off  the  stain,  and  then  stain  2  to  10  minutes  in  the  anilin  blue. 
Dip  in  95  per  cent  alcohol  to  rinse  off  the  stain,  and  treat  for  a  few 
seconds  with  alcohol  slightly  acidulated  with  hydrochloric  acid 
(one  drop  to  50  c.c.  of  95  per  cent  alcohol).  In  the  acid  alcohol 
the  blue  will  become  more  intense,  but  the  red  would  soon  be  ex- 
tracted. Wash  in  95  per  cent  alcohol  to  remove  the  acid.  If  the 
acid  has  weakened  the  Magdala  red,  put  a  pinch  of  sodium  carbonate 
into  the  95  per  cent  alcohol.  The  red  may  brighten.  If  the  red  is 
too  weak,  return  to  the  Magdala  red  and  try  again.  From  the  95 
per  cent  alcohol,  transfer  to  absolute  alcohol,  to  xylol,  and  then 
mount  in  balsam. 

Acid  Fuchsin  and  Iodine  Green  Mixtures. — Two  solutions  are 
kept  separate,  since  they  do  not  retain  their  efficiency  long  after 
they  are  mixed: 

Fuchsin  acid 0 . 1  g. 

Distilled  water 50 . 0  c.c. 

,  Iodine  green 0. 1  g. 

Distilled  water 50 . 0  c.c. 

Absolute  alcohol 100 . 0  c.c. 

Glacial  acetic  acid 1.0  c.c. 

Iodine 0.1  g. 


Stains  and  Staining  63 

Mix  equal  parts  of  A  and  B.  Transfer  to  the  stain  from  water. 
The  proper  time  must  be  determined  by  experiment.  For  a  trial, 
24  hours  might  be  recommended.  Transfer  from  the  stain  directly 
to  solution  C  and  from  C  to  xylol. 

Another  formula: 

A.  Acid  f uchsin 0  5  g. 

B.  Iodine  green 0 . 5  g. 

Mix  a  pipette  full  of  A  with  a  pipette  full  of  B;  stain  2  to  8 
minutes;  transfer  to  85  per  cent  or  95  per  cent,  alcohol,  dehydrate 
rapidly,  clear  in  xylol,  and  mount  in  balsam.  Both  these  formulae 
are  good  for  mitosis. 

Acid  Fuchsin  and  Methyl  Green. — Both  may  be  used  in  1  per 
cent  aqueous-  solutions. 

For  mitotic  figures,  stain  in  green  for  about  an  hour,  wash  in 
water  or  alcohol  until  the  green  is  extracted  from  the  spindle,  and 
then  stain  for  about  one  minute  in  the  fuchsin.  Dehydrate  in  95 
and  100  per  cent  alcohol,  clear  in  xylol  or  clove  oil,  and  mount  in 
balsam.  If  the  green  washes  out,  stain  longer;  if  it  is  not  readily 
extracted  from  the  spindle,  shorten  the  period.  If  the  fuchsin  stains 
the  chromosomes,  shorten  the  period,  and  lengthen  it  if  the  fuchsin 
washes  out  from  the  spindle.  The  chromosomes  should  take  a 
brilliant  green  and  the  spindle  a  bright  red. 

Delafield's  Haematoxylin  and  Erythrosin. — Stain  first  in  the 
haematoxylin,  and  after  that  stain  is  satisfactory,  stain  for  30  seconds 
or  1  minute  in  erythrosin.  This  is  a  good  combination,  and,  for 
most  plant  structures,  gives  a  far  better  differentiation  than  the 
traditional  haematoxylin  and  eosin,  since  the  erythrosin  has  all  the 
advantages  of  the  eosin  and  is  more  transparent.  Orange  G  is  also 
a  good  stain  to  use  with  Delafield's  haematoxylin. 

Directions  for  staining  in  safranin  and  Delafield's  haematoxylin 
are  given  in  the  chapter  on  "Freehand  Sections"  (chap.  vi). 

Haidenhain's  Iron-Haematoxylin  and  Orange  G.— This  haema- 
toxylin is  very  satisfactory  when  used  alone.  A  light  staining  in 
orange  G,  however,  sometimes  improves  the  mount.  After  the  last 
washing  in  water,  stain  for  about  30  seconds  in  orange  G.  Eosin, 
erythrosin,  and  nearly  all  plasma  stains  fail  to  increase  the  effect  of  a 
good  stain  in  iron-haematoxylin. 


64  Methods  in  Plant  Histology 

We  have  not  attempted  to  make  the  list  of  stains  complete.  It  is 
better  to  master  a  few  stains  than  to  use  many  stains  indifferently. 
A  successful  photographer  once  advised  an  amateur  to  stick  to  one 
brand  of  plate  and  one  formula  for  developer.  His  hint  might  well 
have  a  wider  application.  If  one  really  masters  two  or  three  good 
combinations,  he  is  well  prepared  to  develop  methods  for  meeting 
special  needs. 


CHAPTER  IV 
GENERAL  REMARKS  ON  STAINING 

Many  things  may  be. examined  alive  without  killing,  fixing, 
staining,  or  any  of  those  processes.  A  filament  of  Spirogyra  shows 
the  chromatophore  nicely  if  merely  mounted  in  a  drop  of  water; 
the  nucleus  may  be  visible  and  the  pyrenoids  can  usually  be  located. 
Of  course,  such  a  study  is  necessary  if  one  is  to  understand  anything 
about  the  plant,  and  in  an  elementary  class  this  might  be  sufficient, 
but  a  drop  of  iodine  solution  applied  to  the  edge  of  the  cover  would 
emphasize  certain  details,  e.g.,  the  starch  would  appear  blue,  the 
nucleus  a  light  brown,  and  the  cytoplasm  a  lighter  brown.  This 
illustrates  at  least  one  advantage  to  be  gained  by  staining;  it  enables 
us  to  see  structures  which  would  otherwise  be  invisible,  or  almost 
invisible. 

SELECTION  OF  A  STAIN 

With  so  many  stains  at  our  disposal,  it  at  once  becomes  a  problem 
just  which  stain  or  combination  to  use  in  each  particular  case. 
Beautiful  and  instructive  preparations  occasionally  result  from  some 
happy  chance,  but  uniform  success  demands  skill  and  judgment  in 
manipulation,  and  also  a  knowledge  of  the  structures  which  are  to 
be  differentiated.  Let  us  take  a  vascular  bundle  for  illustration. 
Safranin  stains  the  xylem  a  bright  red,  but,  with  judicious  washing, 
is  entirely  removed  from  the  cambium  and  cellulose  elements  of  the 
phloem.  A  careful  staining  with  Delafield's  haematoxylin  now  gives 
a  rich  purple  color  to  the  cellulose  elements  which  were  left  unstained 
by  the  safranin,  thus  contrasting  sharply  with  the  lignified  elements. 
If  cyanin  and  erythrosin  be  used,  the  xylem  takes  the  blue  while  the 
cambium  and  phloem  take  the  red. 

The  mere  selection  of  two  colors  which  contrast  well  is  not 
sufficient.  Green  and  red  contrast  well,  but  safranin  and  iodine 
green  would  be  a  poor  combination,  for  both  would  stain  chromo- 
somes and  neither  would  stain  the  spindle;  both  would  stain  lignified 

65 


66  Methods  in  Plant  Histology 

structures  and  neither  would  give  satisfactory  results  with  cellulose 
walls.  Both  stains  are  basic.  Acid  green  would  have  given  a  con- 
trast in  both  these  cases,  because  it  stains  achromatic  structures  and 
cellulose  walls.  In  general,  an  acid  stain  should  be  combined  with  a 
basic  one,  but  there  are  so  many  exceptions  that  it  is  hardly  worth 
while  to  learn  a  list  of  basic  and  acid  stains.  Stains  which  stain 
chromosomes  are  likely  to  be  basic,  and  those  which  do  not  stain 
chromosomes  are  likely  to  be  acid  or  neutral.  If  it  were  true  that 
acid  stains  affect  only  basic  structures,  and  basic  stains  affect  only 
acid  structures,  a  classification  of  stains  would  be  of  great  value. 
Safranin  and  gentian-violet  are  both  basic,  but  with  proper  washing 
out  the  chromosomes  are  red  and  the  spindle  is  violet,  the  safranin 
being  washed  out  from  the  spindle,  while  the  gentian-violet  is  washed 
out  from  the  chromosomes.  The  only  way  to  insure  success  is  to 
become  familiar  with  the  action  of  each  stain  upon  the  various 
structures. 

THEORIES  OF  STAINING 

In  1890  Auerbach,  a  zoologist,  published  the  results  of  his  studies 
upon  spermatozoa  and  ova.  He  found  that,  if  preparations  contain- 
ing both  spermatozoa  and  ova  were  stained  with  cyanin  and  erythro- 
sin,  the  nuclei  of  the  spermatozoa  took  the  cyanin,  while  the  nuclei 
of  ova  preferred  the  erythrosin;  hence  he  proposed  the  terms  "cya- 
nophilous"  and  "erythrophilous."  Auerbach  regarded  these  differ- 
ences as  an  indication  of  sexual  differences  in  the  cells. 

Rosen  (1892)  supported  this  theory,  and  even  went  so  far  as  to 
regard  the  tube  nucleus  of  the  pollen  grain  as  female,  on  account 
of  its  erythrophilous  staining.  In  connection  with  this  theory  it 
was  suggested  that  the  ordinary  vegetative  nuclei  are  hermaph- 
rodite, and  that  in  the  formation  of  a  female  germ  nucleus  the  male 
elements  are  extruded,  leaving  only  the  erythrophilous  female 
elements;  and,  similarly,  in  the  formation  of  a  male  nucleus  the 
female  elements  are  extruded,  leaving  only  the  cyanophilous  male 
elements. 

As  long  ago  as  1884  Strasburger  discovered  that  with  a  mixture 
of  fuchsin  and  iodine  green  the  generative  nucleus  of  a  pollen  grain 


General  Remarks  on  Staining  67 

stains  green,  while  the  tube  nucleus  stains  red.  In  18921  he  dis- 
cussed quite  thoroughly  the  staining  reactions  of  the  nuclei.  The 
nuclei  of  the  small  prothallial  cells  of  gymnosperm  microspores  are 
cyanophilous  like  the  male  generative  nuclei.  The  nuclei  of  a  nucellus 
surrounding  an  embryo-sac  are  also  cyanophilous,  while  the  nuclei  of 
structures  within  the  sac  are  erythrophilous.  His  conclusion  is  that 
the  cyanophilous  condition  in  both  cases  is  due  to  poor  nutrition, 
while  the  erythrophilous  condition  is  due  to  abundant  nutrition.  A 
further  fact  in  support  of  the  theory  is  that  the  nuclei  of  the  adventi- 
tious embryos  which  come  from  the  nucellus  of  Funkia  ovata  are 
decidedly  erythrophilous,  while  the  nuclei  of  the  nucellus  to  which 
they  owe  their  food-supply  are  cyanophilous. 

In  division  stages  nuclei  are  cyanophilous,  but  from  anaphase 
to  resting  stage  the  cyanophilous  condition  becomes  less  and  less 
pronounced,  and  may  even  gradually  change  to  the  erythrophilous. 

An  additional  fact  in  favor  of  this  theory  is  that  in  Ephedra  the 
tube  nucleus,  which  has  very  little  cytoplasm  about  it,  is  cyanophilous. 
Strasburger  claimed  that  there  is  no  essential  difference  between 
male  and  female  generative  nuclei,  and  subsequent  observation  soon 
showed  that  within  the  oospore  the  sex  nuclei  rapidly  become  alike 
in  their  reaction  to  stains. 

Malfatti  (1891)  and  Lilienfeld  (1892-93)  claim  that  these  reac- 
tions are  dependent  upon  the  amount  of  nucleic  acid  present  in  the 
structures.  During  mitosis  the  chromosomes  consist  of  nearly 
pure  nucleic  acid  and  are  intensely  cyanophilous,  but  the  proto- 
plasm, which  has  little  or  no  nucleic  acid,  is  erythrophilous.  There 
is  a  gradual  transition  from  the  cyanophilous  condition  to  the  ery- 
throphilous, and  vice  versa,  the  acid  structures  taking  basic  stains 
and  basic  structures  the  acid  stains. 

The  terms  " erythrophilous"  and  "cyanophilous"  soon  became 
obsolete,  and  many  claimed  the  affinity  is  for  basic  and  acid  dyes, 
rather  than  for  blue  or  red  colors.  That  the  terms  were  misnomers 
became  evident  when  a  combination  like  safranin  (basic)  and  acid 
green  (acid)  was  used,  for  the  cyanophilous  structures  stained  red, 
and  the  erythrophilous  green. 

^Verhalten  des  Pollens. 


68  Methods  in  Plant  Histology 

According  to  Fischer  (1897  and  1900),  stains  indicate  physical 
but  not  chemical  composition.  Fischer  experimented  with  sub- 
stances of  known  chemical  composition.  Egg  albumin  was  shaken 
until  small  granules  were  secured.  These  were  fixed  with  the  usual 
fixing  agents,  and  then  stained  with  Delafield's  haematoxylin.  The 
extremely  small  granules  stained  red,  while  the  larger  ones  became 
purple.  Since  the  granules  are  all  alike  in  chemical,  composition, 
Fischer  concluded  that  the  difference  in  staining  must  be  due  to 
physical  differences.  With  safranin,  followed  by  gentian-violet, 
the  larger  granules  stain  red  and  the  smaller  violet;  if,  however,  the 
gentian-violet  be  used  first,  then  treated  with  acid  alcohol  and  fol- 
lowed by  safranin,  the  larger  granules  take  the  red  and  the  smaller 
the  gentian-violet.  In  root-tips  similar  results  were  obtained. 
Safranin  followed  by  gentian-violet  stained  chromosomes  red  and 
spindle  fibers  violet,  while  gentian-violet  followed  by  safranin  stained 
the  chromosomes  violet  and  the  spindle  red.  One  often  reads  that 
chromosomes  owe  their  strong  staining  capacity  to  nuclein,  and 
especially  to  the  phosphorous,  but,  according  to  Fischer,  this  is 
shown  to  be  unfounded,  since  albumin  gives  similar  results,  yet 
contains  no  phosphorous,  and  is  not  chemically  allied  to  nuclein. 

Probably  the  most  important  reason  which  led  Fischer  to  under- 
take this  series  of  experiments  was  the  claim  that  certain  granules 
of  the  Cyanophyceae  should  be  identified  as  chromatin  because  they 
behaved  like  chromatin  when  stained  with  haematoxylin.  Fischer's 
experiments  not  only  overthrew  this  claim  but  raised  the  question 
whether  staining  reactions  ever  indicate  chemical  composition.  At 
present,  it  would  seem  that,  in  most  cases,  the  staining  indicates  only 
physical  differences.  However,  in  some  cases  there  is  a  chemical 
reaction,  e.g.,  when  material  fixed  in  bichloride  of  mercury  is  stained 
in  carmine,  mercuric  carminate  is  formed. 

It  would  be  very  convenient  if  we  knew  just  how  much  depend- 
ence should  be  placed  upon  staining  reactions  as  a  means  of  analysis. 
If  two  structures  stain  alike  with  Delafield's  haematoxylin,  does  this 
mean  that  they  have  the  same  chemical  composition;  or  if,  on  the 
other  hand,  they  stain  differently,  must  they  necessarily  be  different 
in  their  chemical  composition?  Delafield's  haematoxylin,  when 


General  Remarks  on  Staining  69 

carefully  used,  gives  a  rich  purple  color,  but  a  careful  examination 
will  often  show  that  in  the  same  preparation  some  structures  stain 
purple,  while  others  stain  red.  Does  this  mean  that  the  purple  and 
red  structures  must  have  a  different  chemical  composition  ?  Many 
people  believe  that  structures  which  stain  differently  with  a  given 
stain  must  be  chemically  different,  but  they  readily  agree  that  struc- 
tures which  stain  alike  are  not  necessarily  similar  in  chemical  com- 
position. Chromosomes  of  dividing  nuclei  and  lignified  cell  walls 
stain  alike  with  safranin;  chromosomes  and  cellulose  cell  walls  stain 
much  alike  with  Delafield's  haematoxylin;  but  everyone  recognizes 
that  the  chromosome  is  very  different  in  its  chemical  composition 
from  either  the  cellulose  or  the  lignified  wall. 

However,  in  an  indirect  and  somewhat  uncertain  way,  one  can 
infer  the  nature  of  certain  structures  from  the  staining.  For  instance, 
if  sections  of  various  objects  have  been  stained  with  safranin,  we  may 
draw  the  following  inferences  with  more  or  less  confidence:  if  cells 
hi  the  xylem  region  of  a  vascular  bundle  stain  red,  their  walls  are 
lignified;  if  cortical  cells,  which  may  appear  quite  similar  in  trans- 
verse section,  stain  red,  they  are  likely  to  be  suberized;  if  the  outer 
walls  of  epidermal  cells  stain  red,  they  are  cutinized;  but  if  the  outer 
boundary  of  the  embryo-sac  of  a  gymnosperm  stains  red,  it  is  chitin- 
ized.-  Of  course,  these  inferences  can  be  made  only  because  the 
various  structures  have  been  tested  by  more  accurate  methods. 

Whatever  doubt  or  uncertainty  there  may  be  in  regard  to  theories 
of  staining  or  in  regard  to  the  value  of  stains  as  a  means  of  analysis, 
there  is  no  doubt  that  stains  are  of  the  highest  importance  in  differ- 
entiating structures,  and  in  bringing  out  details  which  would  other- 
wise be  invisible. 

PRACTICAL  HINTS  ON  STAINING 

The  number  of  stains  in  the  catalogs  is  becoming  so  great  that 
it  is  impossible  to  become  proficient  in  the  use  of  all  of  them.  As 
we  have  already  intimated,  it  is  better  to  master  a  few  of  the  most 
valuable  stains  than  to  do  indifferent  work  with  many.  An  experi- 
enced technician  knows  that  it  is  impossible  to  judge  from  a  few 
trials  whether  a  given  stain  or  combination  is  really  valuable  or 


70  Methods  in  Plant  Histology 

not.  As  a  matter  of  fact,  some  of  the  most  valuable  combinations, 
like  Haidenhain's  iron-alum  haematoxylin  and  Flemming's  safranin, 
gentian-violet,  orange,  require  patient  study  and  long  practice  before 
they  yield  the  magnificent  preparations  of  the  trained  cytologist. 
The  beginner,  especially  if  somewhat  unacquainted  with  the  details 
of  plant  structure,  may  believe  that  he  has  an  excellent  preparation 
when  it  is  really  a  bad,  or  at  most  an  indifferent,  one.  To  illustrate, 
let  us  suppose  that  sections  of  the  pollen  grain  of  a  lily  have  been 
stained  in  safranin  and  gentian-violet.  If  the  preparation  merely 
shows  a  couple  of  dense  nuclei  and  a  mass  of  uniform  cell  contents 
surrounded  by  a  heavy  wall,  the  mount  is  poor.  If  the  two  nuclei 
are  quite  different  and  starch  grains  are  well  differentiated  in  the 
tube  cells  and  the  wall  shows  a  violet  intine  contrasting  sharply 
with  a  red  exine,  the  mount  is  good.  Anything  intermediate  is 
indifferent.  If  mitotic  figures  have  been  stained  with  cyanin  and 
erythrosin,  a  first-class  preparation  should  show  blue  chromosomes 
and  red  spindles;  if  stained  with  safranin  and  gentian-violet,  the 
chromosomes  should  be  red  and  the  spindles  violet. 

In  staining  growing  points,  apical  cells,  young  embryos,  anther- 
idia,  archegonia,  and  many  such  things,  the  cell  walls  are  the  principal 
things  to  be  differentiated,  if  the  preparations  are  for  morphological 
study.  As  a  rule,  it  is  better  in  such  cases  not  to  use  double  stain- 
ing, but  to  select  a  stain  which  stains  the  cell  walls  deeply  without 
obscuring  them  by  staining  starch,  chlorophyll,  and  other  cell  con- 
tents: For  example,  try  the  growing  point  of  Equisetum.  The 
protoplasm  of  such  growing  points  is  very  dense.  If  Delafield's 
haematoxylin  and  erythrosin  be  used,  the  haematoxylin  will  stain 
the  walls  and  nuclei,  and  will  slightly  affect  the  other  cell  contents, 
but  the  erythrosin  will  give  the  cytoplasm  such  a  dense  stain  that 
the  cell  walls  will  be  seriously  obscured.  It  would  be  better  to  use 
haematoxylin  alone.  For  counting  chromosomes,  it  is  better  to  stain 
in  iron-alum  haematoxylin  alone,  or  in  safranin  alone.  The  same 
suggestion  may  well  be  observed  in  tracing  the  development  of 
antheridia,  archegonia,  embryos,  and  similar  structures. 

In  using  combinations,  it  must  be  remembered  that  the  second 
stain  often  affects  the  first,  e.g.,  if  safranin  is  to  be  followed  by 


General  Remarks  on  Staining  71 

Delafield's  haematoxylin  in  staining  a  vascular  bundle,  it  will  not 
do  to  make  the  safranin  just  right  and  then  apply  the  haematoxylin, 
for  the  acid  which  must  be  used  to  differentiate  the  haematoxylin 
and  to  avoid  precipitates  will  also  reduce  the  safranin,  and  the  red 
will  be  too  weak.  You  must  overstain  in  safranin  so  that  the  re- 
duction will  finally  leave  it  just  right.  The  same  hint  will  apply  if 
safranin  is  to  be  followed  by  anilin  blue,  since  here,  also,  acid  must  be 
used;  but  if  light  green  is  to  follow  the  safranin,  no  acid  is  necessary 
and  the  safranin  may  be  made  about  right  before  the  second  stain  is 
added.  These  hints  are  only  samples :  the  student  must  observe  the 
behavior  of  the  various  stains  when  used  singly  and  when  used  in 
various  combinations. 

Permanent  preparations  are  an  absolute  necessity  for  the  greater 
part  of  most  advanced  work,  but  let  us  not  imagine  that  we  cannot 
examine  anything  until  we  have  made  a  permanent  mount.  It 
would  be  impossible  to  make  a  permanent  mount  of  the  rotation  of 
protoplasm.  It  is  better  for  many  purposes  to  look  at  motile  spores 
while  they  are  moving.  Use  Spirogyra  while  it  is  fresh  and  green, 
and  use  permanent  preparations  only  to  bring  out  nuclei  and  other 
details  which  are  not  so  easily  seen  in  living  material.  Examples 
might  be  multiplied. 


CHAPTER  V 
TEMPORARY  MOUNTS  AND  MICROCHEMICAL  TESTS 

Before  the  coming  of  the  microtome  and  the  paraffin  method, 
investigators  were  forced  to  develop  considerable  skill  in  cutting  free- 
hand sections  and  in  teasing  with  needles  and  in  making  delicate 
dissections  under  the  simple  microscope.  Every  student  should 
acquire  some  facility  in  making  mounts  for  immediate  use.  The 
investigator  who  fancies  that  he  cannot  examine  a  structure  until  he 
has  a  carefully  stained  microtome  section  will  not  make  much  prog- 
ress in  modern  botany.  That  particular  class  of  temporary  mounts 
intended  only  for  chemical  tests  is  considered  separately  in  the 
second  part  of  this  chapter. 

TEMPORARY  MOUNTS 

A  preliminary  examination  of  almost  any  botanical  material 
may  be  made  without  any  fixing,  imbedding,  or  staining.  If  a  little 
starch  be  scraped  from  a  potato,  and  a  small  drop  of  water  and  a 
cover-glass  be  added,  a  very  good  view  will  be  obtained,  and  if  a 
small  drop  of  iodine  solution  be  allowed  to  run  under  the  cover,  the 
preparation,  while  it  lasts,  is  better  than  some  permanent  mounts. 
The  unicellular  and  filamentous  algae  can  be  studied  quite  satis- 
factorily from  such  mounts.  The  protonema  of  mosses  and  the 
prothallia  of  ferns  should  be  studied  in  this  way,  even  if  a  later 
study  from  sections  is  intended.  The  addition  of  a  little  iodine 
identifies  the  starch  and  makes  the  nucleus  more  plainly  visible. 
If  the  top  of  a  moss  capsule  be  cut  off  at  the  level  of  the  annulus,  a 
beautiful  view  of  the  peristome  may  be  obtained  by  simply  mounting 
in  a  drop  of  water,  or,  in  a  case  like  this  where  no  collapse  is  to  be 
anticipated,  the  object  may  be  mounted  in  a  small  drop  of  glycerin — 
just  enough  to  come  to  the  edge  of  the  cover  without  oozing  out 
beyond — and  the  preparation  may  be  made  permanent  by  sealing 
with  gold  size  or  any  good  cement.  The  antheridia  and  archegonia 

72 


Temporary  Mounts  and  Microchemical  Tests  73 

of  mosses  may  be  examined  if  the  surrounding  leaves  are  carefully 
teased  away  with  needles.  Freehand  sectioning  with  a  sharp  razor 
and  judicious  teasing  with  a  pair  of  needles  will  give  a  fair  insight  into 
the  anatomy  of  the  higher  plants  without  demanding  any  further 
knowledge  of  technic.  This  rough  work  is  a  'very  desirable  ante- 
cedent to  the  study  of  microtome  sections,  because  most  students  see 
in  a  series  of  microtome 

sections  only  a  series  of    Q^  | 

sections    when,    in    the 

.      .,  FIG.  15.— The  hanging-drop  culture 

mind's  eye,  they  ought 

to  see  the  object  building  itself  up  in  length,  breadth,  and  thickness 

as  they  pass  from  one  section  to  another. 

The  movements  of  protoplasm  can,  of  course,  be  studied  only  in 
the  living  "material.  Every  laboratory  should  keep  Chara  growing 
at  every  season  of  the  year.  Mount  a  small  portion  and  note  the 
movements  in  the  internodal  cells.  Avoid  any  pressure  and  any 
lowering  of  the  temperature.  A  gentle  raising  of  the  temperature 
will  accelerate  the  movements.  Elodea  shows  the  movements  very 
clearly,  especially  in  the  midrib  region.  The  stamen  hairs  of  Trades- 
cantia  have  long  been  used,  their  color,  resembling  a  faint  haema- 
toxylin  stain,  making  them  particularly  favorable.  Stinging  hairs 
show  a  brisk  movement  if  they  are  mounted  quickly  and  without 
injury.  Fortunately,  the  common  onion  always  furnishes  favorable 
material  for  demonstrating  the  movements  of  protoplasm.  Strip 
the  epidermis  from  one  of  the  inner  scales  of  the  bulb  and  mount 

£==============i___mm  m    water.    The 

granu  es  may 
appear  to  better 

FIG.  16. — Another  hanging-drop  culture  , 

advantage    m 
yellow  light,  like  that  of  an  ordinary  kerosene  lamp. 

The  germination  of  spores  and  the  growth  of  pollen  tubes  can 
be  studied  in  the  hanging  drop.  For  facilitating  such  cultures 
there  are  many  devices,  such  as  hollow-ground  slides,  glass  rings, 
rubber  rings,  etc.  (Fig.  15).  A  device  which  is  better  for  most 
purposes,  and  which  is  easily  made  by  any  student,  is  shown  in 
Fig.  16. 


74  Methods  in  Plant  Histology 

A  square  or  round  hole  f  inch  in  diameter  is  cut  in  a  piece  of 
pasteboard  |  inch  thick,  1  inch  wide,  and  1|  inches  long.  The  paste- 
board is  then  boiled  to  sterilize  it  and  to  make  it  fit  more  closely  to 
the  slide.  While  the  pasteboard  is  still  wet,  press  it  to  the  slide, 
make  the  culture  in  a  drop  of  water  or  culture  solution  on  the  cover, 
and  invert  the  cover  over  the  hole.  A  little  water  added  at  the  edge 
of  the  pasteboard  from  time  to  time  will  keep  it  from  warping  and 
will  at  the  same  time  provide  a  constant  moist  chamber. 

In  collecting  material  for  mitotic  figures  in  anthers  it  is  neces- 
sary to  examine  fresh  anthers,  if  one  wishes  to  avoid  a  tedious  and 
uncertain  search  after  the  anthers  have  been  imbedded.  By  teasing 
out  a  few  cells  from  the  apex  and  a  few  from  the  base  of  the  anther 
the  stage  of  development  is  readily  determined,  and  anthers  which 
do  not  show  the  desired  stages  can  be  rejected.  By  allowing  a  drop 
of  eosin  or  methyl  green  to  run  under  the  cover  the  figures  are  more 
easily  detected.  The  actual  progress  of  mitosis  has  been  observed 
in  living  stamen  hairs  of  Tradescantia. 

MICROCHEMICAL  TESTS 

During  the  past  ten  years  there  has  been  such  an  advance  in 
botanical  microchemistry  that  whole  books  are  devoted  to  the 
subject,  and  it  would  be  impossible  to  give  any  complete  presentation 
in  a  book  intended  for  students  of  morphology.  Pflanzenmikro- 
chemie,  by  Dr.  O.  Tunmann  (Gebriider  Borntraeger,  Berlin),  is 
recommended  to  those  who  read  German.  Zimmerman's  Botanical 
Microtechnique  (Henry  Holt  &  Co.,  New  York)  is  still  recommended 
to  those  who  must  rely  upon  English.  We  shall  give  only  a  few 
tests,  but  in  considering  the  various  stains  we  shall  indicate  the 
effect  of  each  stain  upon  the  various  plant  structures. 

Starch. — Mount  the  starch  or  starch-containing  structures  in 
water,  and  allow  a  drop  of  iodine  solution  to  run  under  the  cover. 
Starch  assumes  a  characteristic  blue  color.  The  solution  may  be 
prepared  by  dissolving  1  g.  of  potassium  iodide  in  100  c.c.  of  water 
and  adding  0 . 3  g.  of  sublimed  iodine.  A  strong  solution  of  iodine  in 
alcohol  (about  1  g.  in  50  c.c.  of  absolute  alcohol)  keeps  well.  A  drop 
of  this  solution  added  to  1  c.c.  of  water  is  good  for  testing.  With  too 
strong  a  solution,  the  starch  first  turns  blue  but  rapidly  becomes  black. 


Temporary  Mounts  and  Microchemical  Tests  75 

Grape-Sugar.— In  cells  containing  grape-sugar,  bright-red  gran- 
ules of  cuprous  oxide  are  precipitated  by  Fehling's  solution.  It  is 
better  to  keep  the  three  ingredients  in  separate  bottles,  because  the 
solution  does  not  keep  long  after  they  are  mixed.  The  solutions 
may  be  labeled  A,  B,  and  C. 

f  Cupric  sulphate 3  g. 

A  \Water 100  c.c. 

f  Sodium  potassium  tartrate  (Rochelle  salt) 16  g. 

1  Water 100  c.c. 

{  Caustic  soda 12  g. 

\  Water 100  c.c. 

When  needed  for  use,  add  to  10  c.c.  of  water  5  c.c.  from  each  of 
the  three  solutions.  The  sections,  which  should  be  two  or  three 
cells  in  thickness,  are  warmed  in  the  solution  until  little  bubbles  are 
formed.  Too  much  heat  must  be  avoided.  Mount  and  examine  in 
a  few  drops  of  the  solution.  The  twig  or  organ  may  be  treated  with 
the  solution,  and  the  sections  may  be  cut  afterward.  Other  sub- 
stances precipitate  copper,  and  may  be  mistaken  for  grape-sugar 
by  the  beginner. 

Cane-Sugar. — Cuprous  oxide  is  not  precipitated  from  Fehling's 
solution  by  cane-sugar,  but  after  continued  boiling  in  this  solution 
the  cane-sugar  is  changed  to  invert-sugar  and  the  copper  is  precipi- 
tated. The  solution  becomes  blue. 

Proteids. — The  proteids  turn  yellow  or  brown  with  the  iodine 
solution.  It  is  better  to  use  a  stronger  solution  than  when  testing 
for  starch.  It  must  be  remembered  that  many  other  substances  also 
turn  brown  when  treated  with  iodine. 

When  proteids  are  warmed  gently  in  concentrated  nitric  acid, 
the  acid  becomes  yellow.  The  color  may  be  deepened  by  the  addi- 
tion of  a  little  ammonia  or  caustic  potash. 

When  proteids  are  heated  with  Millon's  reagent,  the  solution 
becomes  brick-red  or  rose-red.  This  reaction  taked  place  slowly 
even  in  the  cold.  The  following  is  one  formula  for  this  reagent: 

Mercury *•  c-c- 

Concentrated  nitric  acid 9  c-c- 

Water 10  c-c- 

Dissolve  the  mercury  in  the  nitric  acid  and  add  the  water. 


76  Methods  in  Plant  Histology 

Fats  and  Oils. — The  fatty  oils  are  not  soluble  in  water  and  are 
only  slightly  soluble  in  ordinary  alcohol.  They  dissolve  readily  in 
chloroform,  ether,  carbon  disulphide,  or  methyl  alcohol. 

Alcannin  colors  oils  and  fats  deep  red.  The  test  is  not  decisive, 
because  ethereal  oils  and  resins  take  the  same  red  color.  Dissolve 
commercial  alcannin  in  absolute  alcohol,  add  an  equal  volume  of 
water,  and  filter.  The  fats  and  oils  in  sections  left  in  this  solution 
for  24  hours  should  be  bright  red.  The  reaction  is  hastened  by 
gentle  heating. 

Osmic  acid,  as  used  in  fixing  agents,  colors  fats  and  oils  brown 
or  black.  The  dark  color  is  removed  by  bleaching  in  a  3  to  10  per 
cent  solution  of  hydrogen  peroxide. 

In  case  of  fats  and  oils,  solubility  and  color  reactions  are  useful, 
but  must  be  regarded  as  corroborative  evidence,  not  as  decisive 
proof.  For  more  critical  and  detailed  methods,  consult  the  book  by 
Tunmann,  which  will  also  give  the  literature  of  the  subject. 

The  Middle  Lamella. — Even  the  origin  and  development  of  the 
middle  lamella  is  none  too  well  known;  its  microchemistry  has 
progressed  but  little  beyond  the  color-reaction  stage.  The  middle 
lamella  consists  largely  of  pectin  or  pectic  compounds.  The  easy 
isolation  of  cells,  when  treated  with  Schultze's  maceration,  depends 
upon  the  ready  solubility  of  pectins  in  this  reagent.  Many  inter- 
cellular spaces  arise  through  the  natural  solution  or  gelatinization 
of  the  lamella. 

In  polarized  light,  with  crossed  Nichols,  the  middle  lamella  is 
resolved  into  three  lamellae,  the  middle  one  appearing  dark,  and 
the  two  outer  lamellae,  light. 

Ruthenium  red  is  a  good  stain,  since  it  gives  as  good  results  as 
any  and  has  the  advantage  of  keeping  well  in  balsam  or  glycerin 
jelly.  Make  a  very  weak  solution— 1  g.  to  5,000  c.c.  of  water, 
or  even  weaker — and  keep  it  in  the  dark.  It  stains  many  other 
things  besides  the  lamella,  but  is,  nevertheless,  a  good  stain. 

Pectin  is  not  at  all  confined  to  the  middle  lamella,  but  is  found 
in  other  membranes,  particularly  in  spore  coats. 

Cellulose. — In  concentrated  sulphuric  acid  cellulose  swells  and 
finally  dissolves.  It  is  also  soluble  in  cuprammonia.  The  cupram- 


Temporary  Mounts  and  Microchemical  Tests  77 

monia  can  be  prepared  by  pouring  15  per  cent  ammonia  water  upon 
copper  turnings  or  filings.  Let  the  solution  stand  in  an  open  bottle. 
It  does  not  keep  well,  but  its  efficiency  is  readily  tested.  Cotton 
dissolves  almost  immediately  as  long  as  the  solution  is  fit  for  use. 

With  iodine  and  sulphuric  acid  cellulose  turns  blue.  Treat 
first  with  the  undiluted  iodine-potassium-iodide  solution  described 
in  the  test  for  starch,  then  add  a  mixture  of  two  parts  of  concentrated 
sulphuric  acid  and  one  part  of  water. 

With  chloroiodide  of  zinc  cellulose  turns  violet.  Dissolve 
commercial  chloroiodide  of  zinc  in  about  its  own  weight  of  water  and 
add  enough  metallic  iodine  to  give  the  solution  a  deep-brown  color. 

The  cell  walls  of  fungi  consist  of  fungus  cellulose.  When  young, 
they  give  a  typical  cellulose  reaction;  when  older,  they  become 
insoluble  in  cuprammonia  and,  with  iodine  and  sulphuric  acid,  show 
only  a  yellow  or  brown,  instead  of  the  typical  blue.  With  chloroiodide 
of  zinc,  the  wall  stains  yellow  or  brown,  instead  of  violet. 

Reserve  cellulose,  which  is  common  in  thick-walled  endosperm  of 
seeds,  shows  the  same  microchemical  reactions  as  ordinary  cellulose. 

Callose. — The  thickening  on  the  sieve  plate  differs  from  cellulose 
in  its  staining  reactions,  and  in  its  solubility.  It  is  insoluble  in 
cuprammonia,  but  will  dissolve  in  a  1  per  cent  solution  of  caustic 
soda. 

Stain  in  a  4  per  cent  aqueous  solution  of  soda  (NaiC02)  for  10 
minutes,  and  transfer  to  glycerin.  The  callus  should  take  a  bright 
red.  If  stained  very  deeply  and  then  transferred  to  a  4  per  cent  soda 
(without  the  corallin),  the  stain  is  extracted  from  the  cellulose  but 
remains  in  the  callus.  Unfortunately,  the  preparations  are  not 
permanent. 

If  stained  for  about  an  hour  in  a  dilute  aqueous  solution  of  anilin 
blue,  the  stain  may  be  extracted  with  glycerin  until  it  remains  only 
in  the  callus.  After  the  blue  is  satisfactory,  a  few  minutes  in  aqueous 
eosin  will  afford  a  good  contrast.  The  preparation  may  be  mounted 
in  balsam  and  is  fairly  permanent. 

Lignin.— Lignified  walls  are  insoluble  in  cuprammonia.  The 
iodine  and  sulphuric  acid  or  the  chloroiodide  of  zinc,  used  as  in  testing 
for  cellulose,  give  the  lignified  walls  a  yellow  or  brown  color.  After 


78  Methods  in  Plant  Histology 

a  treatment  with  Schultze's  maceration  fluid,  lignified  membranes 
react  like  cellulose. 

Phloroglucin  in  a  5  per  cent  aqueous  or  alcoholic  solution  applied 
simultaneously  with  hydrochloric  acid  gives  lignified  walls  a  reddish- 
violet  color.  The  preparations  do  not  keep. 

Cutinized  and  Suberized  Walls. — These  are  insoluble  in  cupram- 
monia  or  concentrated  sulphuric  acid.  They  are  colored  yellow  or 
brown  by  chloroiiodide  of  zinc,  or  by  iodine  and  sulphuric  acid,  when 
applied  as  in  testing  for  cellulose  or  lignin.  With  alcannin,  they 
take  a  red  color,  but  the  red  is  not  as  deep  as  in  case  of  fats  and  oils. 
After  soaking  in  an  aqueous  solution  of  caustic  potash,  suberized 
membranes  take  a  red-violet  color  when  treated  with  chloroi'odide 
of  zinc. 

If  a  strong,  fresh  alcoholic  solution  of  chlorophyll  be  allowed  to 
act  upon  suberized  membranes  for  15  to  30  minutes  in  the  dark, 
they  stain  green,  while  lignified  and  cellulose  walls  do  not  take  the 
stain.  The  preparations  are  not  permanent. 

A  solution  of  alcannin  in  50  per  cent  alcohol  stains  suberized  and 
cutinized  walls  red,  but  the  color  may  not  be  very  sharp. 

Cyanin  can  be  recommended.  First,  treat  with  Eau  de  Javelle 
(potassium  hypochlorite),  which  can  be  obtained  ready  for  use  at 
any  drug-store.  This  destroys  tannins,  and  the  lignified  walls  lose 
their  staining  capacity.  Make  a  1  per  cent  solution  of  cyanin 
(Griibler's)  in  50  per  cent  alcohol  and  add  an  equal  volume  of  glycerin. 
This  should  show  blue  suberized  walls,  while  the  lignified  walls 
remain  unstained. 

Gum,  Mucilage,  and  Gelatinized  Membranes. — These  are  all 
soluble  in  water  and  are  further  characterized  by  their  strong  power 
of  swelling.  They  are  insoluble  in  alcohol.  A  series  of  forms  with 
various  color  reactions  is  included  under  this  heading. 

Crystals. — Nearly  all  crystals  which  are  found  in  plants  consist 
of  calcium  oxalate.  Crystals  of  calcium  carbonate,  calcium  tartrate, 
and  calcium  sulphate  also  occur.  Calcium  oxalate  is  soluble  in 
hydrochloric  acid  or  nitric  acid.  It  is  better  to  use  the  concentrated 
acids.  The  crystals  are  insoluble  in  water  and  acetic  acid.  Sul- 
phuric acid  changes  calcium  oxalate  into  calcium  sulphate.  When 


•  Temporary  Mounts  and  Microchemical  Tests  79 

treated  with  barium  chloride,  crystals  of  calcium  sulphate  become 
covered  with  a  granular  layer  of  barium  sulphate,  while  crystals  of 
calcium  oxalate  are  not  affected. 

Calcium  carbonate,  when  treated  with  hydrochloric  acid  or 
acetic  acid,  dissolves  with  effervescence.  The  acetic  acid  should  be 
rather  dilute. 


CHAPTER  VI 
FREEHAND  SECTIONS 

Sections  which  may  be  cut  without  imbedding,  whether  they  are 
really  cut  freehand  or  with  the  aid  of  a  microtome,  will  be  considered 
here.  The  chapter  will  also  deal  with  other  small  or  thin  objects 
which  may  be  treated  like  freehand  sections. 

The  beginner  is  advised  to  start  with  the  freehand  section,  because 
the  processes  are  rapid,  and  it  is  comparatively  easy  to  find  the  causes 
of  imperfections  and  failures.  In  the  paraffin  method,  where  the 
processes  are  more  complicated,  it  is  often  difficult,  or  even  impossible, 
to  determine  the  exact  cause  of  a  failure. 

As  a  matter  of  fact,  real  freehand  sections,  cut  by  holding  the 
object  in  one  hand  and  the  knife  in  the  other,  are  becoming  less  and 
less  frequent  in  well-equipped  laboratories.  However,  the  laboratory 
is  no  place  for  one  who  is  awkward  with  the  hands;  a  certain  amount 
of  manual  dexterity  must  be  acquired  if  there  is  to  be  any  success  in 
morphological  studies  which  demand  critical  preparations.  Although 
we  know  the  student  will  turn  at  once  to  the  microtome,  we  venture  a 
few  remarks  in  regard  to  real  freehand  sections. 

A  sharp  razor  is  a  necessity.  For  cutting  sections  of  twigs,  roots, 
rhizomes,  and  similar  objects,  a  razor  like  the  one  shown  in  Fig.  7,  A, 
should  be  used;  while  for  sections  of  soft  tissues,  like  young  aspara- 
gus stems,  young  ovaries  of  plants,  most  leaves,  and  such  things,  the 
type  of  razor  shown  in  Fig.  7,  B,  is  much  better.  In  cutting,  brace 
the  forearms  against  the  sides,  hold  the  object  firmly  in  the  left  hand, 
and  cut  with  a  long,  oblique  stroke  from  left  to  right.  The  edge  of  the 
razor  and  the  direction  of  the  stroke  should  be  toward  the  body, 
not  away  from  it  as  in  whittling.  If  the  material  is  fresh,  the  object 
and  the  razor  should  be  kept  wet  with  water,  the  razor  being  dipped 
in  water  for  every  stroke.  For  hard  objects,  like  twigs  of  oak  or 
maple,  the  razor  will  need  sharpening  after  cutting  a  dozen  sections. 
It  is  a  waste  of  time  to  put  off  sharpening  until  the  razor  has  become 


Freehand  Sections  81 

noticeably  dull,  for  all  sections  except  those  cut  when  the  razor  is 
perfectly  sharp  are  sure  to  be  inferior.  With  softer  material  the 
razor  may  hold  its  edge  for  hundreds  of  sections.  Those  sections 
which  seem  to  be  worth  further  treatment  should  be  placed  at  once 
in  water  or  in  a  fixing  agent.1 

With  the  advent  of  a  cheap,  efficient  sliding  microtome,  the 
hand  microtome  began  to  fall  into  disuse  and,  today,  it  has  almost 
disappeared. 

The  sliding  microtome  (Fig.  2)  reduces  to  a  minimum  the  necessity 
for  manual  dexterity,  but  it  is  a  more  complicated  machine.  Study 
the  various  parts  before  you  begin  to  cut  sections.  How  is  the  knife 
adjusted?  How  is  the  object  clamp  raised  and  lowered?  How  is 
the  thickness  of  the  section  determined  ?  In  case  of  a  simple  micro- 
tome like  the  one  shown  in  Fig.  2,  the  student  should  soon  answer 
such  questions  without  any  help  from  the  instructor.  In  case  of 
more  complicated  microtomes,  a  demonstration  by  the  instructor 
will  save  both  time  and  machine. 

In  cutting  sections  of  wood  or  herbaceous-  stems,  the  knife  should 
be  set  obliquely  so  as  to  use  as  much  as  possible  of  the  cutting  edge. 
In  most  cases  it  is  neither  necessary  nor  desirable  to  cut  very  thin 
sections  by  this  method;  10 n  is  very  thin,  and  20,  30,  or  even  40 n 
is  usually  thin  enough. 

Cut  with  a  firm,  even  stroke,  wetting  both  knife  and  object  after 
every  section.  Use  water,  if  the  material  is  fresh;  if  preserved, 
use  the  preservative.  Some  use  a  brush  in  removing  sections  from 
the  knife,  but  nothing  is  quite  equal  to  one's  finger;  anyone  who  is 
in  danger  of  a  cut  while  performing  this  act  is  in  need  of  this  little 
practice  in  manual  dexterity. 

WOODY  AND  HERBACEOUS  SECTIONS 

Safranin  and  Delafield's  Haematoxylin. — In  order  to  make  the 
directions  as  explicit  as  possible,  let  us  follow  the  processes  from 
collecting  the  material  to  labeling  the  slide.  The  rhizome  of  Pteris 
aquilina  is  a  good  object  to  begin  with.  Dig  down  carefully  until 
the  rhizome  is  exposed;  then  with  a  sharp  knife  cut  off  pieces  a  few 

1  See  chap,  ix,  last  three  lines  of  first  paragraph. 


g2  Methods  in  Plant  Histology 

inches  in  length,  taking  the  greatest  care  not  to  strain  the  tissues. 
If  the  rhizome  has  been  cut  carelessly  or  pulled  up,  as  is  usually  the 
case,  the  finished  mount  will  show  ruptures  between  the  bundles  and 
bundle  sheaths,  making  your  work  look  like  the  preparations  sold 
by  optical  companies. 

While  the  material  is  still  fresh  and  moist,  cut  the  sections  and 
place  them  at  once  in  95  per  cent  alcohol,  where  they  should  remain 
20  to  30  minutes.  It  is  not  necessary  to  use  a  large  quantity  of 
alcohol;  10  c.c.  is  enough  for  100  thin  sections  of  the  rhizome. 

Pour  off  the  alcohol  and  pour  on  an  alcoholic  solution  of  safranin 
(a  1  per  cent  solution  of  safranin  in  50  per  cent  alcohol.  See  chap. 
xxix  on  "Formulae  for  Reagents").  It  is  better  to  let  the  safranin 
act  over  night,  or  even  for  24  hours. 

Pour  off  the  safranin  (which  may  be  used  repeatedly)  and  pour 
on  50  per  cent  alcohol.  The  alcohol  will  gradually  wash  out  the 
safranin,  but  this  stain  is  washed  out  more  rapidly  from  cellulose 
walls  than  from  those  which  are  lignified.  The  sections  should 
remain  in  the  alcohol  until  the  stain  is  nearly — but  not  quite — washed 
out  from  the  cellulose  walls,  while  still  showing  a  brilliant  red  in  the 
large  lignified  tracheids.  If  5  or  10  minutes  in  the  alcohol  draws  the 
safranin  from  the  lignified  walls  as  well  as  the  cellulose,  stain  longer; 
if  the  differentiation  is  not  secured  in  5  or  10  minutes,  a  small  drop  of 
hydrochloric  acid  added  to  the  alcohol  will  hasten  the  process. 
Some  recommend  staining  for  only  1  or  2  hours,  but  the  washing-out 
process  is  likely  to  be  rapid  and  uncertain. 

Pour  off  the  alcohol  and  wash  the  sections  thoroughly  in  ordinary 
drinking-water.  The  washing  should  be  particularly  thorough  if 
acid  has  been  used  to  hasten  the  previous  process,  for  the  preparations 
will  fade  if  any  acid  remains. 

Stain  in  Delafield's  haematoxylin  3  to  30  minutes.  Usually  5 
minutes  will  be  about  right.  Delafield's  haematoxylin  will  stain 
the  cellulose  walls,  but  will  have  little  or  no  effect  upon  lignified 
structures. 

Transfer  to  drinking-water,  not  distilled  water.  The  red  color 
of  the  whole  section,  as  it  appears  to  the  naked  eye,  will  be  rapidly 
replaced  by  a  rich  purple.  Continue  to  wash  in  water  for  2  or  3 


Freehand  Sections  83 

minutes  after  the  purple  color  appears.  If  the  cellulose  walls  show 
only  a  faint  purplish  color,  put  the  sections  back  into  the  stain  and 
try  a  longer  period.  If  the  color  is  a  deep  purple  or  nearly  black, 
add  a  little  hydrochloric  acid  (one  drop  to  50  c.c.  is  enough)  to 
the  water.  It  is  better  to  put  the  drop  into  a  bottle  of  water  and 
shake  thoroughly  before  letting  the  acidified  water  act  upon  the 
sections.  As  soon  as  the  sections  begin  to  appear  reddish,  which  may 
be  within  4  or  5  seconds,  pour  off  the  acidified  water  and  wash  in 
drinking-water,  changing  the  water  three  or  four  times  a  minute,  until 
the  reddish  color  caused  by  the  acid  has  been  replaced  by  the  rich 
purple  color  so  characteristic  of  haematoxylin.  The  acid  not  only 
secures  differentiation  by  dissolving  out  the  stain  from  lignified 
structures  more  rapidly  than  from  cellulose  walls,  but  it  also  removes 
the  disfiguring  precipitates  which  almost  invariably  accompany 
staining  with  Delafield's  haematoxylin.  The  acid  also  washes  out 
the  safranin;  it  is  for  this  reason  that  the  washing  after  safranin 
should  be  stopped  while  there  is  still  some  red  color  in  the  cellulose 
walls.  The  acid  should  not  only  reduce  the  density  of  the  haema- 
toxylin and  remove  precipitates,  but  should  also  remove  the  little 
safranin  which  may  remain  in  the  cellulose  walls.  After  the  purple 
color  has  appeared,  the  sections  should  be  left  in  water  for  20  or  30 
minutes.  They  might  be  left  for  several  hours. 

Now  place  the  sections  in  50  per  cent  alcohol  for  1  minute,  then 
in  95  per  cent  alcohol  for  1  minute,  100  per  cent  alcohol  for  5  minutes, 
and  then  transfer  to  xylol.  As  soon  as  the  sections  become  clear — 
in  about  1  to  5  minutes — they  are  ready  for  mounting  in  balsam.  If 
the  sections  do  not  clear  readily,  as  may  be  the  case  if  the  air  is 
damp,  or  if  the  alcohol  or  xylol  is  not  quite  pure,  transfer  from  the 
absolute  alcohol  to  clove  oil,  which  will  clear,  even  if  the  absolute 
alcohol  is  rather  poor.  Then  transfer  from  clove  oil  to  xylol;  the 
objection  to  mounting  directly  from  clove  oil  is  that  preparations 
harden  more  slowly  than  when  mounted  from  xylol.  With  a  section- 
lifter,  or  scalpel,  or  brush,  transfer  three  or  four  sections  to  a  clean, 
dry  slide,  put  on  one  or  two  drops  of  balsam,  and  add  a  cover,  first 
heating  it  gently  to  remove  moisture.  If  xylol  has  been  used  for 
clearing,  it  is  necessary  to  work  rapidly;  for  the  sections  must  never 


81 


Methods  in  Plant  Histology 


be  allowed  to  dry.  Use  square  or  oblong  covers  for  such  mounts, 
reserving  round  covers  for  glycerin  mounts.  If  material  is  abundant, 
use  as  many  sections  as  you  can  cover  conveniently.  If  you  have 
used  several  stains  with  the  same  material,  select  for  each  mount 
sections  from  the  different  stains.  In  ordinary  wood  sections  each 
mount  should  show  the  three  most  important  views,  transverse, 
longitudinal  radial,  and  longitudinal  tangential  sections.  It  is 
wasteful  to  use  three  slides  and  three  covers  to  show  these  three 
views,  or  to  make  a  mount  containing  only  a  single  section  of  the 
rhizome  of  Pteris. 

Put  the  label  at  the  left.     Write  first  the  genus  and  species;  then 
indicate  what  part  of  the  plant  has  been  mounted.     The  date  on 


AXVtowt 

kYv\Axavtv»tt« 


FIG.  17.— The  label 

which  the  material  was  fixed  is  often  valuable.  After  a  year  or  so, 
the  date  of  making  the  mount  may  be  of  interest  in  indicating  the 
relative  durability  of  stains.  The  beginner  is  likely  to  write  also  the 
stains  used,  and  other  details,  which  he  will  find  quite  unnecessary 
after  a  little  experience.  Fig.  17  illustrates  a  good  style  of  labeling 
and  mounting. 

The  following  is  a  convenient  summary  of  the  foregoing  processes, 
beginning  with  the  sections  in  95  per  cent  alcohol : 

1.  Sections  in  95  per  cent  alcohol. 

2.  Safranin,  12  to  24  hours. 

3.  50  per  cent  alcohol,  with  or  without  acid,  until  color  is  right,  gener- 
ally about  2  to  10  minutes. 

4.  Water,  5  minutes,  changing  frequently. 

5.  Delafield's  haematoxylin,  3  to  30  minutes. 

6.  Water,  5  to  10  minutes,  changing  frequently. 

7.  Water  slightly  acidulated,  5  to  10  seconds. 

8.  Water,  to  wash  out  acid,  20  to  30  minutes. 


Freehand  Sections  85 

9.  50  per  cent  alcohol,  1  minute. 

10.  95  per  cent  alcohol,  1  minute. 

11.  100  per  cent  alcohol,  5  minutes. 

12.  Xylol,  1  to  5  minutes. 

13.  Balsam. 

14.  Cover  and  label. 

If  clove  oil  seems  necessary,  finish  as  follows : 

12.  Clove  oil,  2  to  5  minutes. 

13.  Xylol,  1  to  5  minutes. 

14.  Balsam. 

15.  Cover  and  label. 

Since  it  usually  happens  that  processes  are  commenced,  but 
cannot  be  completed  at  a  single  laboratory  period,  it  is  necessary 
to  know  where 'Sections  may  be  left  for  several  hours  or  until  the 
next  day  without  suffering  injury.  At  1,  2,'  or  the  pure  water  of  8  in 
the  schedule  above  given,  sections  may  be  left  until  the  next  day.  If 
it  is  not  desirable  to  mount  all  of  the  sections  which  have  been  pre- 
pared, they  may  be  kept  indefinitely  in  clove  oil  or  xylol.  If  the 
sections  are  to  remain  for  a  year  or  more  in  the  clearing  agent,  xylol 
is  to  be  preferred.  Shells  with  good  corks  are  best  for  keeping  such 
material. 

For  the  study  of  vascular  anatomy,  this  is  the  most  permanent 
stain  which  has  come  into  general  use. 

More  recently,  safranin  combined  with  anilin  blue  or  with 
light  green  has  been  coming  into  favor.  Both  these  methods  will  be 
described. 

Safranin  and  Anilin  Blue. — Use  the  alcoholic  safranin  already 
described,  and  a  1  per  cent  solution  of  anilin  blue  in  90  per  cent 
alcohol. 

With  this  combination  we  should  recommend  a  long  stain  in 
safranin,  not  less  than  24  hours.  Wash  in  50  per  cent  alcohol,  but 
do  not  extract  all  the  safranin  from  the  cellulose  walls.  Stain  2  to  10 
minutes  in  anilin  blue.  Rinse  a  few  seconds  in  95  per  cent  alcohol, 
then  treat  for  about  5  seconds  with  95  per  cent  alcohol  slightly 
acidulated  with  hydrochloric  acid.  The  weak  blue  should  at  once 
change  to  a  bright  blue  and,  at  the  same  time,  the  acid  will  remove 
some  of  the  safranin.  It  is  for  this  reason  that  we  proceed  while 


86  Methods  in  Plant  Histology 

the  sections  are  still  somewhat  overstained  in  safranin.  Wash  for 
1  or  2  minutes  in  95  per  cent  alcohol  to  remove  the  acid.  A  trace  of 
sodium  carbonate,  just  enough  to  make  the  alcohol  alkaline,  may  be 
added  to  the  95  per  cent  alcohol.  If  any  acid  remains,  the  safranin 
will  fade.  Dehydrate  in  absolute  alcohol  1  to  5  minutes,  clear  in 
xylol,  or  first  in  clove  oil  and  then  in  xylol,  and  mount  in  balsam. 

For  convenient  reference,  the  process  may  be  summarized,  but 
it  must  be  remembered  that  all  the  schedules  are  intended  merely  to 
introduce  the  method  to  the  beginner. 

1.  Sections  in  95  per  cent  alcohol. 

2.  Stain  in  safranin,  24  hours. 

3.  50  per  cent  alcohol  until  the  stain  becomes  weak  in  cellulose  walls, 
but  not  until  it  is  removed  entirely. 

4.  Anilin  blue,  2  to  10  minutes. 

5.  95  per  cent  alcohol,  2  to  5  seconds.  , 

6.  95  per  cent  alcohol,  slightly  acidulated  with  hydrochloric  acid, 
5  seconds. 

7.  95  per  cent  alcohol,  with  or  without  a  trace  of  sodium  carbonate, 
1  or  2  minutes. 

8.  Absolute  alcohol,  1  to  5  minutes. 

9.  Xylol,  1  to  5  minutes.    The  xylol  may  be  preceded  by  clove  oil. 
10.  Mount  in  balsam. 

Lignified  and  suberized  walls  should  stain  bright  red  and  cellulose 
walls  bright  blue.  To  make  this  beautiful  combination  a  success,  it 
is  necessary  to  be  very  careful.  If  too  much  safranin  is  extracted 
at  stage  3,  the  acid  at  stage  6  will  still  further  weaken  the  red  stain 
and  the  contrast  will  not  be  sharp. 

Safranin  and  Light  Green  (Land's  Schedule). — This  is  another 
beautiful  combination  and  the  student  should  be  successful  from 
the  first,  since  the  light  green  is  simpler  to  apply  than  either  Dela- 
field's  haematoxylin  or  anilin  blue. 

Land  uses  either  aqueous,  anilin,  or  alcoholic  safranin,  and 
uses  the  light  green  in  clove  oil,  or  in  a  mixture  of  clove  oil  and 
absolute  alcohol.  Make  a  saturated  solution  of  light  green  in  clove 
oil.  Since  the  solution  takes  place  slowly,  the  mixture  should  stand 
several  days  before  using.  If  a  small  quantity  of  absolute  alcohol 
be  added  to  the  clove  oil,  the  stain  dissolves  more  readily.  For 


Freehand  Sections  87 

some  structures  the  stain  is  more  brilliant  than  with  the  simple 
clove-oil  solution. 

Sections  from  fresh  material  are  fixed  in  95  per  cent  alcohol; 
sections  from  preserved  material  are  rinsed  in  alcohol  or  water  before 
staining.  The  following  schedule  will  summarize  the  method: 

1.  Safranin,  2  to  24  hours. 

2.  50  per  cent  alcohol,  until  differentiated. 

3.  Dehydrate  in  95  and  100  per  cent  alcohol. 

4.  Light  green  (in  clove  oil),  3  to  30  minutes. 

5.  Xylol:  2  or  3  c.c.  of  absolute  alcohol  may  be  added  to  each  100  c.c.  of 
xylol,  if  the  free  light  green  shows  a  tendency  to  precipitate. 

6.  Mount  in  balsam. 

This  stain  is  particularly  good  for  phloem.  Since  the  light  green 
is  not  likely  to'  overstain  and  does  not  extract  the  safranin,  the 
combination  is  a  rather  easy  one.  Even  the  beginner  can  hardly 
fail  to  get  a  good  preparation. 

Malachite  Green  and  Congo  Red.— I  am  indebted  to  Dr.  Sharp 
for  this  method,  which  has  been  popular  in  Professor  Gregoire's 
laboratory  at  Lou  vain. 

Sections  of  fresh  material  should  be  treated  with  95  per  cent 
alcohol  and  then  transferred  to  water. 

1.  3  per  cent  aqueous  solution  of  malachite  green  or  methylin  blue, 
6  hours  or  more. 

2.  Wash  in  water. 

3.  Congo  red,  1  per  cent  aqueous  solution,  15  minutes. 

4.  Wash  in  water. 

5.  Rinse  in  80  per  cent  alcohol.    As  soon  as  the  malachite  green  or  anilin 
blue  color  appears  through  the  red,  transfer  quickly  to 

6.  Absolute  alcohol. 

7.  Xylol. 

8.  Balsam. 

Iodine  Green  and  Acid  Fuchsin  is  another  good  combination 
for  such  sections.  The  stain  will  be  particularly  brilliant  if  sections 
from  fresh  material  are  fixed  in  1  per  cent  chromo-acetic  acid  for  10 
to  24  hours;  and  then  washed  for  an  hour  in  water.  Beginning 
with  the  sections  in  water,  the  procedure  is  as  follows: 

Stain  in  aqueous  iodine  green  for  12  to  24  hours.  Then  wash  in 
water  until  the  stain  is  nearly  all  washed  out  from  the  cellulose  walls, 


88  Methods  in  Plant  Histology 

but  is  still  brilliant  in  the  lignified  walls.  If  the  stain  acts  for  too 
short  a  time,  the  washing-out  process  necessary  to  remove  the  stain 
from  the  cellulose  walls  will  leave  only  a  pale-green  color  in  the  ligni- 
fied walls.  Stain  in  aqueous  acid  fuchsin  for  2  to  10  minutes.  This 
should  stain  the  cellulose  walls  sharply,  but  should  not  act  long 
enough  to  affect  the  lignified  tissues.  Pour  off  the  stain  (which  may 
be  used  repeatedly),  and  pour  on  95  per  cent  alcohol,  and  imme- 
diately pour  it  off  and  add  absolute  alcohol.  The  95  per  cent  alcohol 
should  not  act  for  more  than  5  or  10  seconds,  its  only  function  being 
to  save  the  more  expensive  absolute  alcohol.  From  10  to  30  seconds 
will  usually  be  long  enough  for  the  absolute  alcohol.  Too  long  a 
period  in  the  alcohols  will  weaken  the  stain.  Clear  in  xylol  or  clove 
oil,  and  mount  in  balsam. 

If  a  50  or  70  per  cent  alcoholic  solution  of  iodine  green  has  been 
used,  the  stain  should  be  washed  out  in  50  per  cent  alcohol;  otherwise 
the  treatment  is  the  same. 

Methyl  Green  (aqueous  solution)  and  Acid  Fuchsin  is  a  good 
combination,  and  the  student  may  find  it  easier  to  get  a  good  differ- 
entiation than  with  iodine  green.  Follow  the  directions  for  the 
aqueous  iodine  green  and  acid  fuchsin.  It  may  be  necessary  to 
wash  more  rapidly,  since  the-methyl  green  is  easily  extracted. 

Other  Combinations  might  be  suggested,  e.g.,  iodine  green  or 
methyl  green  with  Bismarck  brown,  methyl  green  with  Delafield's 
haematoxylin;  orange  G  might  be  added  after  the  safranin  and 
Delafield's  haematoxylin,  and  various  other  stains  might  be  tried. 
In  double  staining  it  is  usually  best  to  combine  a  basic  with  an  acid 
stain.  Green  and  red  make  a  good  contrast,  but  a  section  stained 
with  iodine  green  and  safranin  would  be  a  failure,  because  both 
stains  would  stain  the  xylem  and  neither  would  stain  the  cellulose. 
Both  stains  are  basic-  Red  lignin  and  green  cellulose  could  be  secured 
by  using  safranin  and  acid  green.  Green  lignin  and  red  cellulose, 
as  already  indicated,  can  be  got  with  iodine  green  and  acid  fuchsin. 

The  Time  Required  for  the  different  processes  varies  greatly, 
and  the  time  required  for  a  subsequent  process  is  often  more  or  less 
dependent  upon  the  time  given  to  processes  which  preceded  it. 
Good  mounts  of  sections  of  the  petiole  of  Nuphar  advena  have  been 
secured  from  material  which  had  been  cut,  fixed,  stained  in  safranin 


Freehand  Sections  89 

and  Delafield's  haematoxylin,  and  mounted  in  balsam,  the  entire 
time  being  less  than  30  minutes.  This  is  an  extreme  case,  and 
nothing  is  gained,  except  time,  and  the  saving  of  time  is  apparent 
rather  than  real,  for  the  histologist  always  has  something  to  do  while 
the  sections  are  in  the  stain. 

Preserved  Material. — If  sections  are  to  be  cut  from  material 
preserved  in  formalin,  the  piece  should  be  washed  in  water,  since 
the  odor  is  annoying  and  the  fumes  are  injurious  to  the  eyes. 

The  sections  are  placed  in  the  stain  from  water.  Sections  from 
alcoholic  material  are  transferred  directly  to  the  stain.  If  the 
material  is  in  a  mixture  of  alcohol  and  glycerin,  the  sections  should 
be  washed  in  water  or  50  per  cent  alcohol  until  the  glycerin  has  been 
removed  before  transferring  to  the  stain. 

Some  material  cuts  well  when  fresh,  but  cuts  with  difficulty  when 
preserved.  On  the  other  hand,  some  material  cuts  well  when  pre- 
served, but  hardly  at  all  when  fresh.  Some  material  which  is  too  soft 
to  cut  when  fresh  can  be  cut  with  ease  after  it  has  been  in  formalin 
alcohol  for  a  week  or  more. 

Very  hard  material,  like  oak,  hickory,  maple,  etc.,  should  be 
boiled  in  water  and  treated  with  hydrofluoric  acid  before  any  section- 
ing is  attempted.  Cut  the  material  into  blocks  suitable  for  sections 
and  boil  in  water  for  several  minutes;  then  transfer  to  cold  water  and, 
after  several  minutes,  repeat  the  boiling.  The  alternate  boiling  and 
cooling,  which  should  be  repeated  several  times,  drives  out  the  air. 
Transfer  to  equal  parts  of  commercial  hydrofluoric  acid  and  water. 
From  1  to  3  weeks  will  be  enough  for  most  woods.  Some  oaks,  ebony, 
apple,  etc.,  may  require  a  longer  time  and  the  acid  may  be  used  pure. 
Wash  thoroughly  in  water  for  a  day  or  two.  Then  leave  in  equal 
parts  of  30  per  cent  alcohol  and  glycerin  for  several  days  before 
cutting.  Material  may  be  left  indefinitely  in  the  mixture  of 
glycerin  and  alcohol. 

OBJECTS  MOUNTED  WITHOUT  SECTIONING 

Fern  Prothallia,  mounted  without  sectioning,  make  very  useful 

preparations.     Select  desirable  stages  and  fix  in  chromo-acetic  acid 

for  10  to  24  hours;  wash  in  water  for  3  or  4  hours,  changing  the  water 

frequently;   stain  in  Delafield's  haematoxylin  for  5  to  30  minutes; 


90  Methods  in  Plant  Hi 

wash  in  slightly  acidulated  water  for  a  few  seconds,  and  then  wash 
thoroughly  in  pure  water.  The  prothallia  must  now  be  brought 
through  a  graded  series  of  alcohols,  15,  35,  50,  70,  85,  95,  and  100  per 
cent  being  sufficiently  close  to  prevent  plasmolysis.  Then  use 
mixtures  of  alcohol  and  xylol,  3  parts  absolute  alcohol  and  1  part 
xylol,  2  parts  alcohol  and  2  parts  xylol,  1  part  alcohol  and  3  parts 
xylol,  and  then  pure  xylol.  Then  bring  the  sections  into  a  mixture 
of  xylol  and  balsam,  using  at  least  10  parts  of  xylol  to  1  of  balsam.  If 
left  in  a  shell,  without  corking,  the  xylol  will  soon  evaporate,  so  that 
in  a  few  days  the  prothallia  may  be  mounted.  Use  the  balsam  in 
which  the  material  has  been  standing,  because  any  other  balsam  may 
have  a  different  concentration.  At  every  step  in  the  process  the 
prothallia  should  be  examined  under  a  microscope,  so  that  any 
plasmolysis  may  be  detected.  If  each  succeeding  step  is  tested  with 
a  single  prothallium,  a  general  disaster  may  be  avoided.  If  plas- 
molysis takes  place,  weaken  the  reagent  and  try  another  prothallium. 
When  a  safe  strength  is  found,  bring  on  the  bulk  of  the  material,  and 
use  the  same  method  with  succeeding  steps.  The  dangerous  places 
are  likely  to  be  the  transfer  from  alcohol  to  xylol  and  the  transfer 
from  xylol  to  balsam.  The  process  is  tedious,  but  the  mounts  are 
very  firm  and  durable.  The  Venetian  turpentine  method  is  less 
tedious,  and,  in  our  opinion,  gives  just  as  good  results. 

Sori  of  Ferns. — Instructive  mounts  of  sori  or  of  individual 
sporangia  may  be  made  without  sectioning.  It  is  better  to  choose 
ferns  with  thin  leaves,  since  leaves  thicker  than  those  of  Asplenium 
thelypteroides  are  likely  to  be  unsatisfactory.  If  this  fern  is  at  hand, 
cut  off  several  of  the  small  lobes  which  bear  three  to  six  pairs  of  sori. 
Fix  in  chromo-acetic  acid;  wash  in  water;  stain  in  Delafield's 
haematoxylin,  or  omit  staining  altogether;  pass  through  a  series  of 
alcohols,  allowing  each  grade  to  act  for  at  least  10  minutes;  clear  in 
clove  oil,  and  mount  in  balsam.  If  the  sori  have  begun  to  turn  brown, 
better  views  of  the  annulus  will  be  obtained  without  staining. 

Mosses  and  Liverworts. — Nearly  all  mounts  are  more  successful 
by  other  methods,  for  which  the  student  should  consult  the  chapters 
on  Bryophytes  (chaps,  xviii,  xix) .  Excellent  mounts  of  the  peristome 
of  the  moss  can  be  made  as  follows:  From  fresh  or  preserved  capsules 
cut  off  the  peristome  just  below  the  annulus.  Treat  with  95  per  cent 


Freehand  Sections  91 

alcohol  1  minute,  absolute  alcohol  2  to  5  minutes,  clear  in  clove  oil  or 
xylol,  and  mount  in  balsam.  It  is  a  good  plan  to  put  at  least  three 
peristomes  on  a  slide,  one  with  the  outside  up,  one  with  the  inside  up, 
and  another  dissected  to  show  details  of  the  teeth. 

Fairly  good  unstained  mounts  of  the  archegonia  and  antheridia 
of  small  mosses  can  be  obtained  by  following  the  directions  for 
mounting  the  sori  of  ferns. 

Beautiful  and  instructive  mounts  of  the  more  delicate  foliose 
Jungermanniaceae  can  be  made  by  staining  lightly  in  Delafield's 
haematoxylin  whole  plants,  or  pieces  as  long  as  can  be  covered  con- 
veniently. The  method  is  that  just  given  for  fern  prothallia.  The 
mount  should  show  both  dorsal  and  ventral  views. 

The  Epidermis  shows  its  best  surface  views  without  sectioning. 
Select  some  form  with  large  stomata,  like  Lilium  or  Tulipa,  strip 
pieces  of  epidermis  from  both  sides  of  the  leaf,  and  place  them  imme- 
diately in  absolute  alcohol  for  1  or  2  minutes.  Stain  in  Delafield's 
haematoxylin;  after  this  stain  is  satisfactory  and  all  acid  has  been 
washed  out,  stain  for  1  or  2  minutes  in  aqueous  eosin,  erythrosin,  or 
acid  fuchsin;  place  directly  into  95  per  cent  alcohol  for  a  few  seconds 
(merely  to  save  the  absolute  alcohol),  then  into  absolute  alcohol  for 
about  30  seconds,  and  then  into  clove  oil.  Mount  in  balsam.  The 
epidermis  is  likely  to  curl  and,  unfortunately,  patience  seems  to  be 
the  only  remedy.  In  mounting,  be  careful  to  get  pieces  from  both 
sides  of  the  leaf,  and  be  sure  that  the  pieces  are  outside  up.  The 
inside  of  the  epidermis  is  usually  more  or  less  rough,  on  account  of 
the  mesophyll  torn  off  with  it.  Sedum  purpurascens  will  show  vari- 
ous stages  in  the  development  of  stomata,  even  in  epidermis  stripped 
from  mature  leaves. 

Other  Objects. — The  cases  just  given  will  suggest  other  objects 
which  might  be  mounted  by  this  method.  In  Part  II  of  this  book 
this  method  is  frequently  recommended  for  mounting  certain  struc- 
tures. Aside  from  the  tediousness  of  the  method,  the  principal 
objection  to  it  is  the  danger  from  plasmolysis. 

Nearly  all  objects  which  used  to  be  mounted  in  balsam  without 
sectioning  are  now  handled  more  successfully  by  the  Venetian  tur- 
pentine method.  Consequently,  the  method  just  described  stands 
just  where  it  was  ten  years  ago. 


CHAPTER  VII 
THE  GLYCERIN  METHOD 

Ten  years  ago  it  was  almost  a  universal  custom  to  mount  fila- 
mentous algae  and  fungi  in  glycerin  or  in  glycerin  jelly.  The  method 
is  simple  and  easily  mastered,  but  preparations  must  be  sealed,  and 
even  when  well  sealed  they  do  not  long  survive  the  ordinary  use 
and  abuse  of  the  laboratory.  The  Venetian  turpentine  method  is 
now  used  for  almost  everything  which  went  into  glycerin  ten  years 
ago.  However,  some  things  are  still  mounted  in  glycerin  or  glycerin 
jelly.  Glycerin  preserves,  to  a  considerable  extent,  the  green  of 
chlorophyll,  a  great  advantage  in  mounts  of  moss  protonema  and  simi- 
lar objects.  Filamentous  forms  may  be  arranged  with  needles  with- 
out much  danger,  since  material  does  not  become  brittle  in  glycerin. 

The  method,  from  fixing  to  mounting,  as  used  in  connection  with 
staining  and  without  staining,  will  now  be  described. 

Stained  Preparations. — The  familiar  Spirogyra  is  a  good  form  to 
begin  with.  Fix  in  chromo-acetic  acid  for  24  to  48  hours.  The 
strength  of  the  fixing  agent  must  be  determined  for  every  collection 
of  material.  If  there  is  in  the  laboratory  a  stock  solution  of  1  per 
cent  chromo-acetic  acid  (1  g.  chromic  acid  and  1  c.c.  glacial  acetic 
acid  to  100  c.c.  of  water),  take  one  part  of  this  stock  solution  and 
add  an  equal  amount  of  water;  then  add  1  c.c.  of  glacial  acetic  acid 
to  each  100  c.c.  of  the  weakened  solution.  Place  a  few  filaments 
hi  the  solution  and,  if  plasmolysis  occurs,  weaken  the  solution  by 
adding  water,  and  then  try  again.  When  a  solution  is  found  which 
causes  no  shrinkage,  fix  the  material  for  24  to  48  hours.  Wash  in 
running  water  over  night,  or  for  24  hours.  If  running  water  is  not 
available,  wash  for  at  least  24  hours,  changing  the  water  frequently. 

The  most  satisfactory  stain  is  Haidenhain's  iron-alum  haema- 
toxylin. 

Treat  for  2  hours  with  a  2  per  cent  aqueous  solution  of  ammonia 
sulphate  of  iron.  Wash  in  water  for  20  minutes  and  then  stain  for 

92 


The  Glycerin  Method  93 

3  to  24  hours  in  a  ^  per  cent  aqueous  solution  of  haematoxylin. 
Wash  again  in  water  for  20  minutes,  and  then  place  the  material  for  a 
second  time  in  the  iron  solution.  The  material  must  now  be  exam- 
ined every  few  minutes,  since  the  iron  solution  extracts  the  stain. 
When  the  stain  is  just  right,  wash  in  water  for  1  to  4  hours.  If  the 
iron  solution  is  not  washed  out  thoroughly,  its  continued  action  will 
cause  the  preparations  to  fade. 

Put  the  material  into  10  per  cent  glycerin  (1  part  glycerin  and  9 
parts  water),  and  then  allow  the  water  to  evaporate  gradually  in  a 
place  as  free  from  dust  as  possible.  Minots,  or  watch  crystals,  are 
good  dishes  for  this  purpose.  The  white  glass  covers  of  "Hazel" 
jars  could  hardly  be  surpassed.  Petri  dishes  are  also  good,  but  rather 
expensive.  When  the  glycerin  has  become  about  as  thick  as  pure 
glycerin,  the  material  is  ready  for  mounting.  A  little  to  the  right  of 
the  center  of  the  slide,  place  a  drop  of  glycerin  in  which  the  material  is 
lying.  In  the  drop  place  a  little  of  the  material,  taking  care  not  to  use 
more  than  can  be  spread  out  without  making  a  confusing  tangle.  Use 
scissors  constantly  so  as  not  to  injure  filaments  by  trying  to  pull  them 
out  from  a  tangle.  There  should  be  just  enough  glycerin  to  come  to 
the  edge  of  the  cover-glass,  but  not  any  more,  for  it  is  impossible  to 
seal  a  mount  if  glycerin  has  oozed  out  beyond  the  cover. 

The  mount  should  now  be  sealed.  Canada  balsam,  various 
asphalts,  cements,  and  glues  have  been  used,  but  the  best  and 
cheapest  of  all  seems  to  be  the  ordinary  flat  varnish,  or  gold  size, 
used  by  painters  in  laying  gold  leaf.  Choose  a  gold  size  of  about  the 
color  of  the  varnish  used  for  ordinary  woodwork.  Mounts  which 
had  been  sealed  with  gold  size  more  than  fifty  years  before  have 
been  exhibited  in  perfect  condition,  but  they  must  have  been  hidden 
away  in  some  museum,  for  a  glycerin  mount  would  never  survive 
fifty  years  of  laboratory  use.  The  gold  size,  as  painters  use  it,  is 
likely  to  be  too  thin  for  sealing  mounts.  Put  some  of  it  in  a  one- 
ounce  bottle  with  a  wide  neck  and  leave  the  cork  out  until  the  gold 
size  thickens  a  little.  Should  it  become  too  thick,  thin  it  with 
turpentine. 

Nothing  but  practice  will  enable  one  to  spin  a  good  ring,  but 
a  good  camel's-hair  brush,  a  good  turntable,  and  a  gold  size  neither 


94  Methods  in  Plant  Histology 

too  thick  nor  too  thin  will  facilitate  matters.  Give  the  turntable  a 
spin,  and  with  the  brush  touch  first  the  slide  about  as  far  out  from  the 
cover  as  you  wish  the  ring  to  extend,  then  gradually  approach  the 
cover.  Dip  the  brush  in  the  gold  size  again,  and  gradually  extend 
the  ring  until  it  is  about  one-sixteenth  of  an  inch  wide  on  the  cover. 
The  touch  must  be  extremely  gentle  or  the  cover  will  be  moved.  Do 
not  try  to  put  on  a  thick  ring  the  first  time,  but  let  a  thin  ring  harden 
for  an  hour  (months  would  do  no  damage),  and  then  a  thicker  ring  can 
be  added  without  any  danger.  Thin  rings  are  too  likely  to  be  broken, 


FIG.  18. — Slide,  natural  size,  showing  size  and  form  of  the  ring 

and  thick  rings  are  in  the  way  if  the  preparation  is  to  be  examined 
with  high  powers.  A  medium  ring  is  best,  and  it  should  consist 
of  two  coats,  for  a  crack  would  seldom  appear  at  the  same  place 
in  both  coats.  A  good  shape  and  thickness  for  a  ring  are  shown 
in  Fig.  18. 

The  following  is  a  summary  of  the  foregoing  processes: 

1.  Fix  in  chromo-acetic  acid,  24  to  48  hours. 

2.  Wash  in  water,  24  hours. 

3.  Iron  solution,  2  hours. 

4.  Wash  in  water,  20  minutes. 

5.  £  per  cent  haematoxylin,  3  to  24  hours. 

6.  Wash  in  water,  20  minutes. 

7.  Iron  solution  until  stain  is  right. 

8.  Wash  in  water,  1  to  4  hours. 

9.  10  per  cent  glycerin. 
10.  Mount  and  seal. 

If  the  material  has  been  fixed  in  formalin,  it  should  be  washed  in 
water  for  5  to  10  minutes  before  staining.  Material  preserved  in  70 
per  cent  alcohol  should  be  placed  successively  in  50  per  cent,  35  per 
cent,  15  per  cent  alcohol,  and  then  in  water,  allowing  each  to  act 
for  15  to  30  minutes  before  being  placed  in  the  stain. 

Mayer's  haem-alum  is  also  a  good  stain  for  filamentous  algae 
and  fungi  which  are  to  be  mounted  in  glycerin.  The  process,  after 
fixing  and  washing  in  water,  is  as  follows: 


The  Glycerin  Method  95 

1.  Transfer  to  the  stain  from  water. 

It  is  seldom  necessary  to  stain  longer  than  10  minutes.  As  a 
rule,  it  is  better  to  dilute  the  stain  (about  1  c.c.  to  10  c.c.  of  distilled 
water)  and  allow  it  to  act  for  10  hours  or  over  night. 

2.  Wash  in  water,  20  minutes. 

3.  10  per  cent  glycerin  until  sufficiently  concentrated. 

4.  Mount  and  seal. 

Eosin  is  a  good  stain  for  many  algae  and  fungi,  when  sharp  out- 
lines rather  than  cell  contents  are  to  be  brought  out.  After  the 
material  has  been  fixed  and  washed  in  water,  stain  in  an  aqueous 
solution  of  eosin  for  12  to  24  hours.  Wash  in  water  until  the  stain 
is  about  right.  Since  subsequent  processes  will  extract  a  little 
more  of  the  stain,  the  washing  in  water  must  stop  a  little  before  the 
desired  differentiation  has  been  secured.  Place  in  1  per  cent  acetic 
acid  for  a  few  minutes  to  fix  the  stain.  Then  place  in  10  per  cent 
glycerin  containing  about  1  per  cent  acetic  acid,  and  allow  the 
glycerin  to  concentrate.  The  acetic  acid  is  to  prevent  the  stain 
from  washing  out.  When  the  glycerin  has  reached  the  proper  con- 
centration, mount  and  seal  as  before. 

The  following  is  a  rapid  method  for  forms  like  Eurotium  and 
Penicillium:  Fix  in  100  per  cent  alcohol  about  2  minutes;  stain 
in  aqueous  eosin  5  minutes;  wash  in  water  about  1  minute;  fix 
in  1  per  cent  acetic  acid  1  minute;  then  mount  directly  in  50  per  cent 
glycerin  to  which  about  1  per  cent  acetic  acid  has  been  added.  It  is 
hardly  worth  while  to  try  this  method  with  forms  which  have  large 
cells ;  they  are  almost  sure  to  collapse.  If  a  form  like  Eurotium  passes 
through  the  earlier  processes  without  danger,  but  collapses  when 
put  into  the  50  per  cent  glycerin,  put  it  into  the  10  per  cent  glycerin 
and  allow  the  glycerin  to  concentrate. 

Mounting  without  Fixing  or  Staining. — It  is  sometimes  desirable 
to  retain  the  natural  color  of  an  object.  The  chlorophyll  green  can 
usually  be  preserved  by  mounting  directly  in  glycerin  without  any 
previous  fixing.  Other  colors  also  are  often  preserved  in  this  way. 
Moss  protonema  make  beautiful  preparations  by  this  method.  If 
possible,  select  protonema  showing  the  very  young  moss  plants.  The 
brown  protonema  and  brown  bulbils  preserve  their  color  perfectly. 
Wash  the  dirt  away  from  the  protonema,  which  is  then  placed  in  10 


96  Methods  in  Plant  Histology 

per  cent  glycerin.     The  brown  or  black  spores  of  fungi  are  readily 
mounted  in  this  way. 

The  method  is  very  useful  when  one  finds  a  single  specimen 
of  Pediastrum,  or  any  small  form  which  would  be  lost  in  the  more 
complicated  processes.  Place  a  large  drop  of  10  per  cent  glycerin 
on  a  slide;  with  a  pipette,  transfer  the  object  to  the  drop,  and  allow 
the  glycerin  to  concentrate.  Then  add  a  cover  and  seal  the  mount. 

GLYCERIN  JELLY 

Glycerin  jelly  is  useful  for  objects  which  are  too  large  to  mount 
in  glycerin  without  making  cells.  With  objects  as  large  as  Volvox 
or  branches  of  Chara,  the  glycerin  is  likely  to  ooze  out  beyond  the 
cover,  making  it  difficult  or  impossible  to  seal  the  mount.  Such 
objects  may  be  mounted  in  glycerin  jelly.  The  material  should  be 
put  into  10  per  cent  glycerin,  which  should  be  allowed  to  concentrate 
until  it  is  as  thick  as  pure  glycerin.  The  bottle  containing  the 
glycerin  jelly  is  then  put  into  warm  water  until  the  jelly  melts. 
No  more  heat  should  be  applied  than  is  really  necessary.  Place  a 
drop  of  the  melted  jelly  on  a  warm  slide,  and  place  on  it  the  material 
to  be  mounted.  Add  a  cover,  and  allow  the  mount  to  cool.  In 
cold  weather,  a  glycerin-jelly  mount  is  safe  without  sealing,  but  in 
summer  the  jelly  may  melt.  It  is  better  to  seal  all  glycerin-jelly 
mounts. 

It  is  a  common  practice  to  put  a  small  piece  of  the  glycerin  jelly 
on  the  slide  and  heat  the  slide  until  the  jelly  melts.  The  only  objec- 
tion is  that  one  may  ruin  his  material  by  putting  it  into  the  drop 
while  it  is  too  hot. 


CHAPTER  VIII 
THE  VENETIAN  TURPENTINE  METHOD 

Twenty  years  ago  Pfeiffer  and  Wellheim1  described  a  method  for 
mounting  fresh-water  algae  in  Venetian  turpentine.  The  method 
received  no  recognition  in  the  United  States  and  did  not  become 
current  in  Europe.  I  made  a  casual  trial  of  the  method  when  pre- 
paring the  first  edition  of  this  book,  but  the  preparations  were  such 
miserable  failures  that  the  process  did  not  seem  worth  mentioning. 
The  method  was  next  brought  to  my  attention  during  a  demonstration 
in  Strasburger's  laboratory  at  Bonn.  He  was  using  preparations  of 
Zygnema  and  Spirogyra,  the  staining  of  which  surpassed  anything  I 
had  ever  seen.  He  remarked  that  it  was  not  worth  while  to  consult 
the  lengthy  article,  because  his  preparations  had  been  made  by 
the  authors  and  no  one  else  had  made  a  success  of  the  method.  How- 
ever, when  I  returned,  I  made  a  careful  study  of  the  process,  and 
finally  learned  to  use  it  successfully.  The  details  as  given  in  this 
paper  were  too  indefinite  for  practical  use,  but,  after  one  has  learned 
the  method,  the  article  can  be  read  with  profit. 

The  great  practical  advantages  of  the  method  are  that  prepara- 
tions are  as  hard  and  durable  as  balsam  mounts,  and  that  a  much 
greater  variety  of  staining  is  possible  than  in  case  of  glycerin  mounts. 

After  fixing  and  washing  in  water,  the  general  outline  of  the 
method  is  as  follows: 

1.  10  per  cent  glycerin  until  concentrated. 

2.  Wash  the  glycerin  out  thoroughly  in  95  per  cent  alcohol. 

3.  Stain.     Use  stains  dissolved  in  about  90  per  cent  alcohol. 

4.  Wash  in  95  per  cent  alcohol,  and  complete  the  dehydration  in  100 
per  cent  alcohol. 

5.  10  per  cent  Venetian  turpentine2  in  an  exsiccator  until  the  turpentine 
becomes  thick  enough  for  mounting. 

6.  Mount  in  the  Venetian  turpentine. 

'  Pfeiffer,  Ferdinand,  and  Wellheim,  R.  v.,  "Zur  Preparation  der  Stisswasseralgen," 
Jahrbucher  filr  wissenschaftliche  Botanik,  26:674-732,  1894. 

2  The  Venetian  turpentine  which  we  have  used  is  marked  "Venice  Turpentine 
(true)."  It  can  be  obtained  from  Morrison,  Plummer  &  Co.,  Chicago,  Illinois. 

97 


gg  Methods  in  Plant  Histology 

While  this  is  the  general  outline,  it  is  not  sufficiently  definite  for 
a  working  introduction.  The  following  concrete  examples,  describ- 
ing the  use  of  Venetian  turpentine  with  an  aqueous  stain,  with  an 
alcoholic  stain,  and  with  a  combination  of  aqueous  and  alcoholic 
stains,  will  be  more  practical  than  general  directions.  The  steps 
from  fixing  to  mounting,  as  used  with  an  aqueous  stain,  will  be 
described  first,  since  this  will  introduce  the  method  in  its  least  com- 
plicated form. 

Haidenhain's  Iron-Haematoxylin. — Using  Spirogyra  as  a  type, 
proceed  as  follows: 

1.  Fix  24  hours  in  chromo-acetic  acid. 

1  per  cent  chromic  acid 70  c.c. 

Glacial  acetic  acid 3  c.c. 

Water 90  c.c. 

The  volume  of  the  fixing  agent  should  be  at  least  100  times  that  of 
the  material  to  be  fixed. 

2.  Wash  in  water,  24  hours. 

3.  2  per  cent  aqueous  solution  of  ammonia  sulphate  of  iron,  2  hours. 

4.  Wash  in  running  water,  20  minutes.     If  running  water  is  not  avail- 
able, wash  in  a  large  quantity  of  water  and  change  frequently. 

5.  Stain  over  night,  or  24  hours,   in  £  per   cent  aqueous  solution 
haematoxylin. 

6.  Wash  in  water,  20  minutes. 

7.  2  per  cent  aqueous  solution  of  ammonia  sulphate  of  iron,  until  the 
stain  is  satisfactory.     This  can  be  determined  only  by  examining 
frequently  under  the  microscope. 

8.  Wash  in  water,  2  hours.    If  this  washing  is  not  thorough,  the  con- 
tinued action  of  the  iron-alum  will  cause  the  preparations  to  fade. 

9.  Transfer  to  10  per  cent  glycerin,  and  allow  the  glycerin  to  concentrate 
until  it  has  the  consistency  of  pure  glycerin.     It  is  not  necessary  to 
use  an  exsiccator.     Merely  put  the  glycerin  into  shallow  dishes,  and 
leave  it  exposed  to  the  air,  but  protected  from  dust.     If  the  material 
is  in  Petri  dishes  or  other  dishes  with  a  large  surface,  3  or  4  days 
will  be  sufficient.    This  process  should  not  be  hastened  by  warming. 

10.  Wash  out  the  glycerin  with  95  per  cent  alcohol.    It  will  be  necessary 
to  change  the  alcohol  several  times.     From  10  to  20  minutes  will  be 
sufficient  if  the  alcohol  is  changed  frequently. 

11.  Complete  the  dehydration  in  100  per  cent  alcohol:    10  minutes 
should  be  sufficient. 

12.  Most  failures  are  now  ready  to  occur. 


The  Venetian  Turpentine  Method  99 

From  the  absolute  alcohol  the  material  is  transferred  to  a  10 
per  cent  solution  of  Venetian  turpentine  in  absolute  alcohol.  The 
turpentine  thickens  as  the  alcohol  evaporates,  and  when  it  reaches  the 
consistency  of  pure  glycerin  the  material  is  ready  for  mounting. 
The  10  per  cent  Venetian  turpentine  is  very  sensitive  to  moisture,  and 
most  failures  are  due  to  this  characteristic;  consequently  the  con- 
centration cannot  be  allowed  to  take  place  with  the  turpentine 
exposed  to  the  air  of  the  room.  Use  an  exsiccator.  This  will  not 
only  absorb  the  moisture  from  the  air,  but  will  soon  remove  the 
alcohol  from  the  turpentine  mixture.  Make  an  exsiccator  as  follows: 
Place  a  saucer  full  of  soda  lime  (sodium  hydroxide  with  lime)  on  a 
plate  of  glass,  and  cover  with  a  bell  jar.  This  is  a  simple  and  effect- 
ive exsiccator.  Instead,  you  may  simply  scatter  soda  lime  in  the 
bottom  of  any  low  museum  jar  with  tight-fitting  cover.  The 
saucer  of  soda  lime  may  be  placed  on  a  smooth  board  and  covered 
with  a  perfectly  tight  box.  You  may  improvise  other  forms;  the 
essential  thing  is  to  provide  a  small  air-tight  space  in  which  the  soda 
lime  may  work. 

Instead  of  soda  lime  you  may  use  fused  calcium  chloride  or  the 
white  sticks  of  sodium  hydroxide. 

Paint  the  exsiccator  black,  or  cover  it  with  black  paper,  or  in 
some  other  way  shut  out  the  light.  Many  stains  are  weakened 
by  light. 

We  are  now  ready  for  the  transfer  from  absolute  alcohol  to  the 
10  per  cent  Venetian  turpentine.  Make  the  transfer  quickly.  Pour 
off  the  absolute  alcohol  and  place  the  dish,  with  the  material,  in  the 
exsiccator;  then  pour  on  the  10  per  cent  turpentine,  and  immediately 
put  on  the  cover.  This  is  better  than  to  pour  on  the  turpentine  and 
then  try  to  get  the  dish  well  placed  in  the  exsiccator. 

The  greater  the  surface  of  soda  lime  exposed,  the  more  rapid 
will  be  the  concentration  of  the  Venetian  turpentine.  The  con- 
centration must  not  be  too  rapid.  Not  less  than  2  days  should  be 
allowed  for  the  concentration  of  30  c.c.  of  the  turpentine  in  an 
ordinary  Minot  watch  glass. 

Great  care  must  be  taken  not  to  let  any  of  the  soda  lime,  or  other 
drier,  get  into  the  turpentine. 


100  Methods  in  Plant  Histology 

As  soon  as  the  turpentine  has  attained  the  consistency  of  pure 
glycerin,  it  may  be  exposed  to  the  air  without  any  danger  from 
moisture;  but  the  turpentine  would  soon  become  too  thick  for 
mounting.  If  the  turpentine  has  become  too  thick,  thin  it  with 
a  few  drops  of  absolute  alcohol  or  with  10  per  cent  or  any  thin 
solution  of  Venetian  turpentine. 

Mount  the  material  in  a  few  drops  of  the  Venetian  turpentine 
and  add  a  cover.  Square  covers  may  be  used,  since  it  is  entirely 
unnecessary  to  seal  the  mounts.  Such  mounts  are  as  hard  and 
durable  as  balsam  mounts. 

Material  in  the  thickened  Venetian  turpentine,  if  not  needed 
for  immediate  mounting,  may  be  put  into  small  vials  or  shells,  where 
it  can  be  kept  indefinitely.  The  shells  should  be  kept  out  of  the  light. 

We  recommend  a  No.  4  shell.  The  corks  should  be  of  the  best 
quality;  otherwise  the  turpentine  will  become  too  thick.  While  it 
can  be  thinned  by  adding  thin  turpentine,  it  is  better,  for  easy 
mounting,  not  to  let  the  turpentine  become  too  thick. 

Magdala  Red  and  Anilin  Blue. — Fix  in  chromo-acetic  acid  and 
wash  in  water,  as  described  in  the  previous  schedule.  Transfer 
from  water  to  10  per  cent  glycerin  and  allow  the  glycerin  to  con- 
centrate. It  is  not  necessary  to  use  an  exsiccator  since  there  is  no 
danger  from  moisture  in  the  air.  When  the  glycerin  attains  the 
consistency  of  pure  glycerin,  wash  the  glycerin  out  with  95  per  cent 
alcohol  and  then  proceed  with  the  staining. 

1.  Stain  in  Magdala  red.     At  least  two  Magdala  reds  are  sold 
by  dealers.     The  one  marked  edit  is  more  expensive,  but,  in  our 
experience,   is  inferior  to  the  one  marked  simply  Magdala  red. 
Make  a  1  per  cent  solution  in  90  per  cent  alcohol.     We  use  the  stain 
much  stronger  than  recommended  by  Pfeiffer  and  Wellheim.     This 
solution,  diluted  with  an  equal  volume  of  95  per  cent  alcohol  and 
allowed  to  act  for  24  hours,  does  not  stain  too  deeply. 

2.  Rinse  the  material  for  a  minute  in  90  per  cent  alcohol. 

3.  Stain  in  anilin  blue,  using  a  1  per  cent  solution  in  90  per  cent 
alcohol,  diluted  with  four  times  its  volume  of  90  per  cent  alcohol.     We 
prefer  to  make  a  fresh  solution  every  time  we  have  anything  to  stain. 
It  is  not  necessary  to  measure  it.     A  little  of  the  powder — about 


The  Venetian  Turpentine  Method  101 

alf  the  bulk  of  a  grain  of  wheat — in  30  c.c.  of  90  per  cent  alcohol, 

all  give  an  efficient  solution.     The  time  required  for  successful 

, taming  will  vary  from  3  to  30  minutes.     Do  not  put  all  the  material 

into  the  anilin  blue  at  once,  but,  by  trying  a  few  filaments  at  a  time, 

find  out  what  the  probable  periods  may  be. 

4.  Rinse  off  the  stain  in  90  per  cent  alcohol,  and  then  treat  for  a 
few'seconds  in  acid  alcohol  (1  very  small  drop  of  HC1  to  30  c.c.  of  90 
oer  cent  alcohol).     The  acid  alcohol  fixes  and  brightens  the  anilin 

lue,  but  extracts  the  Magdala  red.  If  the  anilin  blue  or  the  acid 
alcohol  acts  for  too  short  a  time,  the  blue  will  be  weak;  if  they  act 
too  long,  the  red  is  lost  entirely.  If  the  blue  overstains  too  much, 
wash  it  out  in  95  per  cent  alcohol.  If  the  red  overstains,  wait  until 
the  mount  is  finished,  and  then  reduce  the  red  by  exposing  the  slide 
to  direct  sunlight. 

5.  Absolute  alcohol,  5  or  6  seconds. 

6.  Transfer  quickly  to  10  per  cent  Venetian  turpentine  and  pro- 
ceed as  in  the  previous  schedule. 

The  surprising  beauty  of  successful  preparations  will  compen- 
sate for  whatever  failures  may  occur.  Nuclei  and  pyrenoids  should 
show  a  brilliant  red,  while  the  chromatophores  and  cytoplasm  should 
be  dark  blue.  The  cell  walls  should  show  a  faint  bluish  color. 

Haidenhain's  Iron-Alum  Haematoxylin  and  Eosin.— Follow 
the  schedule  for  iron-haematoxylin  until  the  glycerin  has  been  washed 
out  in  95  per  cent  alcohol.  Then  stain  for  a  minute  in  a  solution  of 
eosin  in  95  per  cent  alcohol.  Wash  for  a  minute  in  95  per  cent  alcohol, 
then  a  minute  in  absolute  alcohol,  and  then  transfer  to  the  10  per  cent 
Venetian  turpentine. 

Other  Stains  may  be  used.  Aqueous  stains  should  be  used  before 
starting  with  the  10  per  cent  glycerin.  Alcoholic  stains  should  be  in 
strong  alcohol — about  90  per  cent — -and  should  be  applied  just  after 
washing  out  the  glycerin. 

This  method  is  equally  good  for  filamentous  fungi  and  also  for 
the  prothallia  of  Equisetum  and  ferns,  for  delicate  liverworts  and 
mosses,  and  similar  objects. 


CHAPTER  IX 
THE  PARAFFIN  METHOD 

The  paraffin  method  is  still  the  most  important  of  all  histological 
methods  now  in  use.  The  results  obtained  by  this  method  would 
have  been  regarded  as  almost  miraculous  by  the  histologists  of  one 
hundred  years  ago.  At  that  time  it  was  customary  to  observe 
things  dry,  and  no  cover-glasses  were  used.  Section-cutting  with 
sharp  knives  or  razors  did  not  become  general  until  about  1830.  The 
need  for  an  instrument  which  would  cut  sections  without  demanding 
an  extreme  degree  of  manual  dexterity  was  soon  felt,  but  a  successful 
microtome  did  not  appear  until  much  later.  The  latest  microtomes, 
while  rather  complicated,  give  wonderful  results.  The  Spencer 
microtome,  shown  in  Fig.  19,  with  the  cooling  attachment  devised 
by  Dr.  Land,  will  cut  even  ribbons,  1  p.  in  thickness,  from  such 
material  as  the  antheridial  receptacles  of  Marchantia.  This  means 
that  a  series  of  sections  can  be  cut  from  pollen  grains  or  spores  too 
small  to  be  seen  by  the  naked  eye.  Many  of  the  principles  involved 
in  this  method  are  general  in  their  application,  and  some  of  the  pro- 
cesses are  common  to  other  methods.  Before  attempting  the  free- 
hand sectioning,  the  beginner  should  read  the  following  paragraphs 
on  killing  and  fixing,  washing,  hardening  and  dehydrating,  and  on 
clearing. 

KILLING  AND  FIXING 

As  stated  in  the  chapter  on  " Reagents"  (chap,  ii),  the 
purpose  of  a  killing  agent  is  to  bring  the  life-processes  to  a  sudden 
termination,  while  a  fixing  agent  is  used  to  fix  the  cells  and  their 
contents  in  as  nearly  the  living  condition  as  possible.  The  fixing 
consists  in  so  hardening  the  material  that  the  various  elements  may 
retain  their  natural  condition  during  all  the  processes  which  are  to 
follow.  Usually  the  same  reagent  is  used  for  both  killing  and  fixing. 
Zoologists,  from  humane  motives,  may  use  chloroform  for  killing, 
while  other  reagents  are  used  for  fixing.  In  fixing  root-tips,  anthers, 

102 


The  Paraffin  Method 


103 


and  other  material  for  a  study  of  mitotic  figures,  it  is  necessary  that 
killing  be  very  prompt.  In  a  weak  solution  of  chromo-acetic  acid, 
nuclei  which  have  begun  to  divide  may  complete  the  division, 
although  the  reagent  might  hinder  nuclei  from  entering  upon  division. 
By  treating  for  20  minutes  to  1  hour  with  Flemming's  weaker 


FIG.  19. — Spencer  rotary  microtome  with  electric  motor  and  Land's  apparatus  for 
temperature  control. 

solution,  or  with  a  chromo-acetic  solution  containing  a  much  smaller 
proportion  of  osmic  acid,  the  killing  will  be  greatly  accelerated  and 
the  proportion  of  nuclei  in  division  will  be  correspondingly  greater. 
If  filamentous  algae  are  placed  for  10  or  20  minutes  in  a  chromo- 
acetic  solution  containing  a  little  osmic  acid,  all  the  advantages  of 
immediate  killing  will  be  secured.  Material  is  then  transferred  to 
chromo-acetic  acid  containing  no  osmic  acid.  The  short  treat- 
ment with  an  osmic  solution  is  not  likely  to  cause  any  serious 
blackening. 


104   '  Methods  in  Plant  Histology 

Take  the  killing  and  fixing  fluids  into  the  field.  If  one  waits 
until  the  material  is  brought  to  the  laboratory  there  may  be  some 
fixing,  but  it  will,  in  many  cases,  be  too  late  to  do  much  killing. 
Material  which  has  begun  to  wilt  is  not  worth  fixing.  Material  like 
Spirogyra,  however,  may  be  brought  from  the  field  into  the  laboratory 
before  fixing,  if  considerable  water  be  brought  with  it.  Branches  with 
developing  buds  may  be  brought  in  and  kept  in  water.  Cones  of  the 
cycad,  Ceratozamia,  sent  from  Jalapa,  Mexico,  have  arrived  in 
Chicago  with  cell  division  still  going  on  at  a  rapid  rate.  But  such 
cases  are  extremes;  as  a  rule,  take  the  killing  and  fixing  fluids  into  the 
field. 

Always  have  the  material  in  very  small  pieces,  in  order  that 
the  reagents  may  act  quickly  on  all  parts  of  the  specimens.  Pieces 
larger  than  cubes  of  1  cm.  should  be  avoided  whenever  possible. 
While  one  sometimes  needs  sections  2  or  even  3  cm.  long,  it  is  not 
likely  to  be  necessary  to  fix  pieces  more  than  4  or  5  mm.  in  thickness. 
For  very  fine  work  no  part  of  the  specimens  should  require  the 
reagent  to  penetrate  more  than  1  or  2  mm. 

For  fixing  agents  of  the  chromic-acid  series,  the  volume  of  the 
reagent  should  be  about  fifty  times  that  of  the  material. 

Fixing  agents  with  alcohol  as  an  ingredient  will  fix  a  larger  pro- 
portion of  material.  It  must  be  remembered  that  the  water,  which 
is  always  present  in  living  tissues,  weakens  the  fixing  agent. 

The  time  required  for  fixing  varies  with  the  reagent,  the  character 
of  the  tissue,  and  the  size  of  the  piece.  About  24  hours  is  a  com- 
monly recommended  period  for  chromic-acid  solutions,  but  2  or  even 
3  days  will  do  no  harm. 

Directions  for  making  and  using  the  various  fixing  agents  are 
given  in  the  chapters  on  "Reagents"  (chaps,  ii,  xxix). 

WASHING 

Nearly  all  fixing  agents,  except  the  alcohols,  must  be  washed  out 
from  the  material  as  completely  as  possible  before  any  further  steps 
are  taken,  because  some  reagents  leave  annoying  precipitates  which 
must  be  removed,  and  others  interfere  with  subsequent  processes. 
Aqueous  fixing  agents  with  chromic  acid  as  their  principal  ingredient 


The  Paraffin  Method  105 

are  washed  out  with  water;  aqueous  solutions  of  corrosive  sublimate 
are  also  washed  out  with  water;  but  alcoholic  solutions  should  be 
washed  out  with  alcohol  of  about  the  same  strength  as  the  fixing 
agent;  picric  acid,  or  fixing  agents  with  picric  acid  as  an  ingredient, 
must  not  be  washed  out  with  water,  but  with  alcohol,  whether  the 
picric  acid  be  in  aqueous  or  alcoholic  solution.  When  washing  with 
water,  running  water  is  best,  and  where  this  is  not  convenient  the 
water  should  at  least  be  changed  frequently.  The  washing-out 
process  usually  requires  about  24  hours. 

HARDENING  AND  DEHYDRATING 

After  the  material  has  been  washed,  it  is  necessary  to  continue 
the  hardening  and  also  to  remove  the  water.  Alcohol  is  used  almost 
entirely  for  these  purposes.  It  completes  the  hardening  and  at  the 
same  time  dehydrates,  that  is,  it  replaces  the  water  in  the  material, 
an  extremely  important  consideration,  for  the  least  trace  of  moisture 
is  likely  to  interfere  seriously  with  the  infiltration  of  the  paraffin. 

The  process  of  hardening  and  dehydrating  must  be  gradual; 
if  the  material  should  be  transferred  directly  from  water  to  absolute 
alcohol,  the  hardening  and  dehydrating  would  be  brought  about 
in  a  very  short  time,  but  the  violent  osmosis  would  cause  a  ruinous 
contraction  of  the  more  delicate  parts.  In  recent  years,  cytologists 
have  been  making  the  dehydration  process  more  and  more  gradual. 
Ten  years  ago  most  workers  began  the  dehydration  process  with 
35  per  cent  alcohol  and  used  the  series  35,  50,  70,  85,  95,  and  100  per 
cent  alcohol.  Some  placed  an  intermediate  grade  between  water  and 
35  per  cent  alcohol.  If  plasmolysis — the  tearing  away  of  the  proto- 
plast from  the  cell  wall — was  avoided,  the  series  was  thought  to  be 
sufficiently  gradual;  but  a  series  which  may  avoid  plasmolysis  may 
not  be  adequate  if  one  is  to  study  the  finer  details  of  cell  structure. 
The  following  series  is  recommended:  2|,  5,  7£,  10,  15,  20,  30,  40,  50, 
70,  85,  95,  and  100  per  cent.  There  is  no  particular  virtue  in  the 
fractions:  it  is  convenient  to  make  10  per  cent  alcohol,  dilute  with 
an  equal  volume  of  water  for  the  5  per  cent,  and  dilute  the  5  per  cent 
with  an  equal  volume  for  the  2|  per  cent.  It  will  be  noted  that  the 
series  begins  with  very  close  grades  and  that  the  intervals  are 


106  Methods  in  Plant  Histology 

gradually  increased.  The  claim  is  that  by  beginning  with  very  weak 
alcohols  in  close  grades,  more  perfect  dehydration  can  be  secured 
at  the  end  of  the  series.  Various  devices,  like  constant  drip  and 
osmotic  apparatus,  have  been  proposed  to  secure  a  more  gradual 
transfer,  but  it  is  very  doubtful  whether  these  possess  any  real  advan- 
tages. It  is  not  necessary  to  use  a  large  amount  of  alcohol:  2  or  3 
times  the  volume  of  the  material  is  sufficient. 

The  grades  of  alcohol  may  be  used  several  times,  but  it  must  be 
remembered  that  pollen  grains,  fungus  spores,  starch  grains,  and  vari- 
ous granules  are  likely  to  be  left  in  the  alcohol,  so  that,  when  it  is 
necessary  to  know  the  identity  of  every  such  structure,  only  pure 
alcohols  should  be  used. 

As  the  alcohols  absorb  water  from  the  material,  they  become 
weaker  and  weaker.  If  the  various  alcohols  be  poured  in  a  large 
"waste  alcohol"  bottle,  when  a  couple  of  liters  has  been  accumulated, 
the  strength  may  be  determined  by  testing  with  an  alcoholometer. 
Then  any  grade  of  less  strength  can  be  made  from  this  stock. 

The  time  necessary  for  each  of  the  stages  has  not  been  determined 
with  any  certainty.  About  4  hours  seems  to  be  long  enough  for 
each  of  the  grades  from  1\  to  70  per  cent;  for  70,  85,  and  95,  about 
10  hours  each;  for  absolute  alcohol,  12  to  24  hours,  changing  two 
or  three  times.  If  material  is  to  be  kept  in  alcohol,  leave  it  in  85  per 
cent,  but  where  labor  is  no  object,  it  is  better  to  go  on  and  imbed 
it  in  paraffin. 

CLEARING 

Let  us  suppose  that  the  material  has  been  thoroughly  dehydrated, 
so  that  not  the  slightest  trace  of  water  remains.  If  the  supposition 
chances  to  be  contrary  to  fact,  all  the  work  which  has  preceded,  as 
well  as  all  which  is  to  follow,  is  only  an  idle  waste  of  time.  The 
purpose  of  a  clearing  agent  is  to  make  the  tissues  transparent,  but 
clearing  agents  also  replace  the  alcohol.  At  this  stage  the  latter 
process  is  the  essential  one,  the  clearing  which  accompanies  it  being 
incidental.  The  clearing,  however,  is  very  convenient,  since  it 
shows  that  the  alcohol  has  been  replaced  and  that  the  material  is 
ready  for  the  next  step. 


The  Paraffin  Method  107 

Various  clearing  agents  are  in  use.  Xylol  is  the  most  generally 
employed,  and  for  most  purposes  it  seems  to  be  the  best.  Bergamot 
oil,  cedar  oil,  clove  oil,  turpentine,  and  chloroform  are  used  for  the 
same  purpose.  Cedar  oil  and  chloroform  may,  in  some  cases,  be  as 
good  as  xylol. 

Only  a  small  quantity  of  the  clearing  agent  is  necessary,  enough 
to  cover  the  material  being  sufficient. 

The  transfer  from  absolute  alcohol  to  the  clearing  agent  should 
be  gradual,  like  the  hardening  and  dehydrating  processes.  The  most 
successful  workers  have  been  making  this  transfer  more  and  more 
gradual.  Twenty  years  ago  it  was  customary  to  transfer  from 
absolute  alcohol  directly  to  xylol;  then  a  mixture  of  equal  parts  of 
absolute  alcohol  and  xylol  was  interpolated;  in  the  second  edition 
of  this  book  three  grades  were  placed  between  the  absolute  alcohol 
and  xylol.  It  is  undoubtedly  better  to  make  the  transfer  still  more 
gradual.  The  following  series  seems  to  be  safe:  2£,  5,  10,  15,  25,  50, 
75,  and  100  per  cent  xylol.  These  mixtures  of  absolute  alcohol  and 
xylol  can  be  made  with  sufficient  accuracy  without  measuring  in  a 
graduate.  The  50  per  cent  grade  is  made  by  mixing  equal  parts  of 
absolute  alcohol  and  xylol;  the  25  per  cent,  by  adding  to  the  50  per 
cent  an  equal  volume  of  absolute  alcohol;  make  the  10  per  cent  grade 
from  the  25  per  cent  by  adding  a  little  more  than  an  equal  volume 
of  absolute  alcohol;  in  the  same  way,  make  the  5  per  cent  from  the 
10  per  cent,  and  the  2£  per  cent  from  the  5  per  cent.  The  different 
grades  may  be  kept  in  bottles  and  may  be  used  repeatedly. 

About  3  or  4  hours  is  enough  for  each  grade.  The  pure  xylol 
should  be  changed  once  or  twice.  Throughout  the  dehydrating  and 
clearing  it  is  a  good  plan  to  keep  the  material  in  No.  4  shells,  which 
are  made  from  glass  tubing  about  25  mm.  in  diameter. 

Other  clearing  agents  may  be  used,  but  the  process  must  be  just 
as  gradual. 

THE  TRANSFER  FROM  CLEARING  AGENT  TO  PARAFFIN 
This  should  also  be  a  gradual  process.    The  most  convenient 
method  is  to  place  a  small  block  of  paraffin  in  the  pure  clearing  agent 
with  the  material,  but  the  block  of  paraffin  should  not  rest  directly 


108  Methods  in  Plant  Histology 

upon  the  objects.  Dr.  Land  uses  coarse  wire  gauze,  cut  into  strips 
about  15  mm.  wide  and  tapered  at  both  ends.  The  strip  is  then  bent 
so  that  the  pointed  ends  rest  upon  the  bottom  of  the  No.  4  shell, 
while  the  middle  portion  forms  a  flat  table  upon  which  the  paraffin 
may  rest.  Dip  the  wire  gauze  table  into  xylol  and  then  slip  it  care- 
fully into  the  No.  4  shell.  The  table  portion  should  be  10  or  15  mm. 
above  the  material,  and  there  should  be  enough  xylol  to  extend  a  few 
millimeters  above  the  table.  Place  on  the  table  a  block  of  paraffin 
about  equal  to  the  volume  of  the  xylol  in  the  shell.  The  table  not 
only  prevents  the  paraffin  from  injuring  the  material  by  mechanical 
pressure,  but  insures  considerable  diffusion  before  the  mixture  of 
paraffin  and  xylol  reaches  the  specimens.  After  24  hours  (or  several 
days,  if  time  permits)  at  room  temperature,  place  the  shell  on  a  thin 
piece  of  wood  or  corrugated  paper  on  the  top  of  the  paraffin  bath. 
Do  not  place  the  shell  directly  upon  the  metal  of  the  bath,  since  it 
is  better  to  minimize  heat.  As  soon  as  the  paraffin  is  dissolved,  add 
some  more,  this  time  leaving  the  cork  out,  in  order  that  the  xylol 
may  evaporate.  About  24  hours  on  the  top  of  the  bath  should  be 

sufficient. 

THE  PARAFFIN  BATH 

This  step  is  usually  called  infiltration,  but  when  the  transfer 
from  the  clearing  fluid  to  paraffin  is  made  gradually,  as  has  just  been 
indicated,  the  process  of  infiltration  is  already  begun.  It  is  now 
necessary  to  get  rid  of  the  xylol  or  other  clearing  agent.  This  id 
accomplished,  to  a  considerable  extent,  by  pouring  off  the  mixture  of 
xylol  and  paraffin  and  replacing  it  with  pure  melted  paraffin;  but 
some  xylol  remains  in  the  tissues  and  must  be  removed.  Do  not  put 
the  shell  into  the  bath,  but  use  a  flat  dish  of  some  sort.  The  main 
object  is  to  have  a  fairly  large  surface  exposed,  so  that  the  remaining 
xylol  may  evaporate  as  rapidly  as  possible.  Change  the  paraffin 
two  or  three  times.  Soft  paraffin  (about  45°  C.)  may  be  used  at 
first,  but  the  second  should  be  the  paraffin  of  the  grade  in  which  the 
material  is  to  be  imbedded.  If  there  are  two  baths,  one  should  be 
kept  at  46°  C.  and  the  other  at  53°  C.,  if  the  material  is  to  be  imbedded 
in  52°  C.  paraffin.  While  using  the  soft  paraffin,  keep  the  material 
in  the  46°  C.  bath;  for  the  harder  paraffin,  use  the  53°  C.  bath. 


The  Paraffin  Method  109 

Do  not  throw  away  the  paraffin  which  you  pour  off,  but  put  it  in  a 
waste  jar  or  beaker,  or,  still  better,  in  a  small  tin  lard  pail,  in  which 
you  have  made  a  lip  to  facilitate  pouring.  This  can  be  placed  in  the 
bath,  or,  in  winter,  on  the  radiator,  and  the  xylol  will  gradually 
evaporate.  After  long  heating,  the  paraffin  not  only  becomes  as 
good  as  new,  but  even  better,  since  it  becomes  more  homogeneous 
and  tenacious.  If  it  contains  dust  or  de"bris  of  any  kind,  it  may  be 
filtered  with  a  hot  filter. 

The  time  required  varies  with  the  character  of  the  material  and 
the  thoroughness  of  the  dehydrating  and  clearing.  If  this  schedule 
has  been  followed  up  to  this  point,  the  time  will  be  much  shorter  than 
most  investigators  now  deem  necessary.  Fern  prothallia  infiltrate 
perfectly  in  15  to  20  minutes;  onion  root-tips  in  20  to  30  minutes; 
ovaries  of  Lilium  at  the  fertilization  stage,  30  to  40  minutes;  5  or 
6  mm.  cubes  of  endosperm  of  cycads,  containing  archegonia,  1  hour 
to  1^  hours;  median  longitudinal  sections,  4  or  5  mm.  thick,  through 
ovulate  cones  of  Pinus  Banksiana  may  require  6  or  8  hours;  if  serial 
sections  through  the  entire  cone  are  wanted,  Miss  Aase  found  that 
the  time  must  be  prolonged  to  24  or  even  48  hours.  When  one  is 
dealing  with  many  lots  of  the  same  kind  of  material,  as  in  research 
work,  the  time  required  for  infiltration  is  easily  determined.  As  a 
rule,  minimize  heat.  It  is,  probably,  never  necessary  to  use  paraffin 
with  a  melting-point  higher  than  52°  C.  With  Land's  cooling  device 
sections  1  /JL  in  thickness  can  be  cut  from  52°  C.  paraffin;  and  sections 
2  or  3  fj,  in  thickness  can  be  cut  from  45°  C.  paraffin. 

IMBEDDING 

Material  may  be  imbedded  in  paper  trays,  Petri  dishes,  watch 
crystals,  or  in  apparatus  made  for  the  purpose.  Many  use  imbedding 
L's  consisting  of  two  L-shaped  pieces  of  brass  or  type  metal.  A  pair 
of  L-shaped  pieces  with  arms  three  inches  long  will  furnish  a  box  of 
almost  any  required  size.  A  piece  of  glass  serves  for  a  bottom.  The 
most  satisfactory  imbedding-dish  we  have  used  is  a  thin  rectangular 
porcelain  dish,  glazed  inside.  This  dish,  called  a  Verbrennungsschale, 
is  made  by  the  Konigliche  Porzellan-Manufactur,  Berlin,  Germany. 
The  most  convenient  sizes  are  40X50X10  mm.,  68X45X10  mm., 


110 


Methods  in  Plant  Histology 


and  91X58X15  mm.  These  are  listed  respectively  at  50,  60,  and 
80  Pfennige.  As  listed,  these  dishes  are  not  glazed;  care  should 
be  taken  to  indicate  that  the  dishes  must  be  glazed  inside  (innen 
glasirt).  The  tray,  Petri  dish,  or  whatever  is  used,  should  be  slightly 
smeared  with  glycerin,  to  prevent  sticking.  If  several  objects 
are  to  be  imbedded  in  one  dish,  it  is  best  to  have  the  dish  as  near  the 


temperature  of  melted  paraffin  as 


IHHMMM 
IIMIflMlf 
tflHIMIff 


otherwise  the  objects 
may  stick  to  the  bottom, 
and  it  will  be  impossible  to 
arrange  them  properly.  Hot 
needles  are  good  for 
arranging  material.  Great 
care  should  be  taken  not  to 
have  the  dish  too  hot,  since 
too  high  a  temperature  not 
only  injures  the  material, 
but  also  prevents  a  thorough 
imbedding.  Pour  the  par- 
affin with  the  objects  into 
the  imbedding  dish  and 
arrange  them  so  as  to  facili- 
tate the  future  cutting  out 
from  the  paraffin  cake. 
Look  at  Figs.  20  and  21, 
representing  the  arrange- 

. 

ment    of  root-tips  in   a  par- 
afgn     cake        From     a     Cake 

like  that  in  Fig.  20  it  is 
easy  to  cut  out  tips  for  sectioning.  The  arrangement,  or  rather  the 
lack  of  it,  shown  in  Fig.  21,  should  be  remembered  only  as  an 
exasperating  example. 

After  the  objects  have  been  arranged,  cool  the  cake  rapidly  by 
allowing  the  bottom  of  the  dish  to  rest  upon  cold  water.  As  soon  as 
a  sufficiently  firm  film  forms  on  the  surface  of  the  cake,  let  water 
flow  gently  over  the  top.  After  the  cake  has  been  under  water  for  a 
minute,  it  may  be  placed  under  the  cold-water  tap  to  complete  the 


Fio.  21 

Fios.  20,  21. — Paraffin  cakes  of  root-tips,  the 
upper  (Fig.  20)  showing  a  good  arrangement,  the 
lower  (Fig.  21)  showing  fewer  tips  and  most  of 
these  not  in  position  to  be  blocked  without  injury. 


The  Paraffin  Method  1U 

cooling.  If  paraffin  cools  slowly  it  crystallizes  and  does  not  cut  well. 
The  layer  of  paraffin  should  be  just  thick  enough  to  cover  the  objects, 
not  only  as  a  matter  of  economy,  but  because  a  thick  layer  retards 
the  cooling.  Very  small  objects,  like  the  megaspores  of  Marsilea, 
ovules  of  Silphium,  etc.,  may  simply  be  poured  out  upon  a  cool  piece 
of  glass.  In  this  way  very  thin  cakes  are  made,  which  harden  very 
rapidly. 

CUTTING 

As  soon  as  the  paraffin  is  thoroughly  cooled,  it  is  ready  for  cutting. 
Trim  the  paraffin  containing  the  object  into  a  convenient  shape,  and 
fasten  it  upon  a  block  of  wood.  Blocks  of  pine  f  inch  long  and  f  inch 
square  are  good  for  general  purposes.  Put  paraffin  on  the  end  of  the 
block  so  as  to  form  a  firm  cap  about  f  inch  thick.  Warm  the  cap 
and  the  bottom  of  the  piece  containing  the  object,  and  press  them 
lightly  together;  then  touch  the  joint  with  a  hot  needle,  put  the 
whole  thing  into  cold  water  for  a  minute,  and  it  is  ready  for  cutting. 
Cutting  can  be  learned  only  by  experience,  but  a  few  hints  may  not 
come  amiss: 

a)  Keep  the  knife  sharp.  There  should  be  two  hones,  one  for 
use  when  the  knife  is  rather  dull  and  the  other  for  finishing.  For 
the  first  hone,  nothing  equals  a  fine  carborundum  hone.  About 
5.5X22.5  cm.  is  a  good  size.  A  hard  Belgian  hone,  of  the  same 
size,  may  be  a  little  better  for  finishing.  Flood  the  stone  with  water, 
and  rub  it  with  the  small  slip  which  accompanies  all  high-grade  hones; 
this  not  only  makes  a  lather  which  facilitates  the  sharpening,  but 
it  also  keeps  the  surface  of  the  hone  flat.  As  soon  as  the  edge  of  the 
knife  appears  smooth  and  even  under  a  magnification  of  thirty  or 
forty  diameters,  the  sharpening  is  completed  with  a  good  strop.  It 
is  better  to  sharpen  the  knife  every  time  you  use  it.  A  first-class 
microtome  knife,  in  perfect  condition,  is  unsurpassed  for  cutting 
paraffin  sections,  but  it  requires  both  time  and  skill  to  keep  the  edge 
perfect.  More  than  ten  years  ago  we  began  to  experiment  with  the 
Gillette  safety-razor  blade  and  devised  several  holders  for  it,  some 
of  them  more  or  less  successful.  Mr.  Strickler  finally  perfected  a 
holder  which  has  already  been  mentioned.  In  using  this  holder 
the  blade  should  not  project  more  than  1  mm. 


112 


Methods  in  Plant  Histology 


6)  Keep  the  microtome  well  oiled  and  clean. 

c)  Trim  the  block  so  that  each  section  shall  be  a  perfect  rectangle. 

A  ribbon  of  sections  like  that  shown  in  Fig.  22,  A,  is  much  better 
than  one  like  B  of  the  same  figure,  because  sections  will  usually  come 
off  in  neater  ribbons  if  the  knife  strikes  the  longer  edge  of  the  rec- 
tangle, so  that  the  sections  are  united  by  the  longer  sides  rather 


FIG.  22.— The  ribbon      , 

than  by  the  shorter.  Crooked  ribbons  are  caused  by  wedge-shaped 
sections,  and  are  always  to  be  avoided,  because  they  make  it  difficult 
to  economize  space,  and  also  because  they  present  such  a  disorderly 
appearance.  The  knife,  which  should 
I  \  be  placed  at  a  right  angle  to  the  block 
and  not  obliquely,  should  strike  the 
whole  edge  of  the  block  at  once,  and 
should  leave  in  the  same  manner. 

If  sections  stick  to  the  knife,  it 
may  be  that  the  knife  is  too  nearly 
parallel  with  the  surface  of  the  block, 
as  in  Fig.  23,  A.  By  inclining  the 
knife  as  in  Fig.  23,  B,  this  difficulty 
is  often  obviated.  In  using  the  safety- 
razor  blade  in  a  handle,  it  must  be  remembered  that  the  blade  is 
sharpened  from  both  sides;  the  angle  must  be  sufficient  to  let  the 
paraffin  block  clear.  A  split  or  scratch  in  the  paraffin  ribbon  may 
be  caused  by  a  nick  in  the  knife.  Use  some  more  favorable 
position  of  the  edge,  or  sharpen  the  whole  knife.  A  split  or  a 
scratch  in  the  ribbon  is  often  caused  by  some  hard  granule  which 
becomes  fastened  to  the  inner  side  of  the  edge  of  the  knife.  This 


FIG.  23.— Position  of  the  knife 


The  Paraffin  Method  113 

is  the  most  common  cause  of  the  difficulty.  Simply  wipe  the  knife 
by  an  upward  stroke  of  the  finger,  slightly  moistened  with  xylol.  Do 
not  use  a  cloth. 

Sometimes  good  sections  can  be  cut  with  a  rather  slow  stroke 
when  a  rapid  stroke  fails.  When  paraffin  is  rather  hard,  sections 
may  sometimes  cut  nicely  at  5  n,  when,  at  10  n,  ribbons  cannot 
be  secured.  If  very  thin  sections  are  desired  and  the  paraffin  seems 
too  soft,  cool  the  paraffin  and  the  edge  of  the  knife  with  ice,  or  better 
still,  by  Land's  cooling  device.  Sometimes  hard  paraffin  does  not 
ribbon  well.  This  difficulty  may  be  remedied  by  dipping  a  hot 
needle  in  soft  paraffin  and  applying  it  to  the  opposite  edges  of  the 
block  to  be  cut.  Often  the  mere  warming  of  the  opposite  edges  of 
the  block  with  a  hot  needle  is  sufficient. 

Another  method,  suggested  by  Dr.  Land  to  facilitate  the  cutting 
of  difficult  material,  has  been  tested  in  this  laboratory  and  has  been 
found  to  be  very  effective.  Paraffin  absorbs  a  small  amount  of 
water,  or  water  penetrates  between  the  crystals  of  paraffin.  At 
any  rate,  water  reaches  cell  walls  and,  perhaps,  other  structures 
which  have  not  been  completely  infiltrated  and  thus  softens  them. 
The  paraffin  cakes  may  be  left  for  weeks  in  water.  Cakes  of  class 
material  may  be  put  in  water  in  a  fruit  can  and  kept  until  ready 
for  use.  After  such  treatment,  smooth  ribbons  may  be  cut  from 
material  which  would  hardly  cut  at  all  without  it. 

FIXING  SECTIONS  TO  THE  SLIDE 

Mayer's  Fixative. — Sections  must  be  firmly  fixed  to  the  slide, 
or  they  will  be  washed  off  during  the  processes  involved  in  staining. 
Mayer's  albumen  fixative  is  excellent  for  this  purpose.  Formula: 

White  of  egg  (active  principle) 50  c.c. 

Glycerin  (to  keep  it  from  drying  up) 50  c.c. 

Salycilate  of  soda  (antiseptic,  to  keep  out  bac- 
teria, etc.) 1  g- 

Shake  well  and  filter.  It  will  keep  from  2  to  6  months,  but,  to  say  the 
least,  it  is  never  better  than  when  first  made  up.  Of  course,  white 
of  egg  may  be  used  alone,  since  the  other  two  ingredients  are  merely 
incidental.  Put  a  small  drop  of  fixative  on  the  slide,  smear  it  evenly 


114  Methods  in  Plant  Histology 

over  the  surface,  and  then  wipe  it  off  with  a  clean  finger  until  only  a 
scarcely  perceptible  film  remains;  then  add  several  drops  of  distilled 
water  and  float  the  sections  or  ribbons  on  the  water.  Warm  gently 
until  the  paraffin  becomes  smooth  and  free  from  wrinkles.  Be  care- 
ful not  to  melt  the  paraffin,  for  the  albumen  of  the  fixative  coagulates 
with  less  heat  than  is  required  to  melt  the  paraffin.  If  the  paraffin 
should  melt,  run  some  cold  water  under  it,  and  transfer  the  ribbon 
to  another  slide,  prepared  with  fixative  and  water.  It  is  a  very  good 
plan  to  put  the  slide  on  a  metal  bath  or  warming  plate,  like  that 
shown  in  Fig.  10.  After  the  sections  have  become  smooth,  remove 
the  surplus  water  and  leave  them  on  the  bath  with  a  couple  of  thick- 
nesses of  blotting  paper  under  them  for  3  or  4  hours,  or,  better,  over 
night.  If  the  fixative  is  used  alone,  as  is  often  the  case  when  sections 
are  very  thick,  none  of  this  delay  is  necessary,  since  the  sections 
are  merely  laid  upon  the  fixative  and  pressed  down  gently  with  the 
finger. 

Land's  Fixative. — Mayer's  fixative  is  so  easily  prepared  and 
it  keeps  so  well  that  it  is  in  universal  use;  but,  in  many  cases,  it  will 
not  hold  the  section  to  the  slide.  Moss  archegonia  and  moss  capsules 
are  likely  to  wash  off,  especially  if  cut  rather  thick.  Large  sections 
of  cones  of  conifers  are  almost  sure  to  float  off  as  soon  as  the  slide 
comes  into  the  xylol  or  alcohol.  Sections  of  ovules  of  cycads,  as 
soon  as  they  attain  a  length  of  1 .5  to  2  cm.,  are  likely  to  wash  off. 
For  handling  these  more  difficult  cases,  Dr.  Land  devised  a  fixative 
which  has  proved  satisfactory,  even  in  such  extreme  cases  as  sections 
of  ovulate  cones  of  Pinus  Banksiana  2  cm.  long.  Formula: 

Gum  arabic 1 . 0  g. 

Bichromate  of  potash 0 . 2  g. 

Water 100.0  c.c. 

The  mixture  will  not  keep;  the  formula  is  given  merely  to  indicate 
its  composition.  Make  a  1  per  cent  solution  of  gum  arabic  in  water, 
which  will  keep  as  well  as  Mayer's  fixative;  but  make  the  bichromate 
solution  immediately  before  using.  Do  not  make  the  solution 
stronger  than  1  per  cent;  usually  0.2  per  cent  is  strong  enough. 
Dr.  Land  does  not  measure,  but  simply  adds  enough  bichromate 
crystals  to  make  the  water  pale  yellow. 


The  Paraffin  Method  115 

Smear  a  few  drops  of  the  1  per  cent  solution  of  gum  arable  on  the 
slide;  flood  with  the  bichromate  solution;  warm  to  straighten  the 
ribbons;  drain  off  the  excess  water  and  let  the  preparation  dry  in 
the  light.  The  exposure  to  light  renders  the  gum  insoluble  in  water. 
LePage's  glue  or  Mayer's  albumen  fixative  may  be  used  instead  of 
gum  arabic. 

The  foregoing  directions  are  taken  from  Dr.  Land's  notes. 

REMOVAL  OF  THE  PARAFFIN 

To  remove  the  paraffin,  place  the  slide  in  a  Stender  dish  of  xylol. 
About  5  minutes  will  be  sufficient  for  sections  10  n  thick.  The  time 
may  be  shortened  a  little  by  gently  warming  the  slide.  Never  heat 
the  slide  enough  to  melt  the  paraffin.  Never  attempt  to  warm  the  paraffin 
over  a  lamp.  Overheating  is  ruinous. 

Many  prefer  to  remove  the  paraffin  by  pouring  on  xylol  or 
turpentine.  Hold  the  slide  at  an  angle  of  45°,  and  pour  on  a 
little  xylol  or  turpentine.  If  the  slide  has  been  slightly  warmed, 
this  should  carry  off  the  paraffin  immediately.  The  reagent  used 
in  this  first  pouring  cannot  be  used  again.  Now  flood  the  slide 
several  times  with  the  turpentine  or  xylol,  pouring  the  reagent 
back  into  the  bottle. 

REMOVAL  OF  XYLOL  OR  TURPENTINE 

To  remove  the  xylol,  place  the  slide  in  equal  parts  of  xylol 
and  absolute  alcohol  in  a  Stender  dish.  After  5  minutes,  trans- 
fer to  absolute  alcohol,  which  should  also  be  allowed  to  act  for 
5  minutes. 

If  the  pouring  process  is  preferred  and  turpentine  has  been  used 
to  remove  the  paraffin,  remove  the  turpentine  by  flooding  the  slide 
with  95  per  cent  alcohol.  About  100  c.c.  of  turpentine  and  200  c.c. 
of  95  per  cent  alcohol  should  be  sufficient  for  fifty  slides,  even  if  the 
sections  are  to  be  mounted  under  the  longest  covers.  By  keeping 
both  reagents  in  bottles  and  pouring  the  liquid  on  the  slide,  the 
reagents  are  always  fresh.  A  given  quantity  of  the  reagent  will 
prepare  as  many  slides  by  one  method  as  by  the  other. 


116  Methods  in  Plant  Histology 

TRANSFER  TO  THE  STAIN 

After  the  paraffin  has  been  removed  with  xylol  or  turpentine,  and 
the  xylol  or  turpentine  has  been  rinsed  off  with  alcohol,  the  next 
step  is  the  staining.  If  the  stain  is  a  strong  alcoholic  one  (85  to  100 
per  cent  alcohol),  transfer  directly  to  the  stain.  If  the  stain  is  in 
70  per  cent  alcohol,  pass  through  95  and  85  per  cent  alcohol,  5  minutes 
in  each,  before  staining.  If  an  aqueous  stain  is  to  be  used,  pass  down 
the  whole  series — 95,  85,  70,  50,  35,  and  water — 5  minutes  in  each, 
before  placing  the  slide  in  the  stain. 

This  is  rather  tedious,  but,  for  cytological  work,  it  seems  to  be 
necessary.  For  general  morphological  work,  the  slide  may  be  trans- 
ferred directly  from  the  absolute  or  95  per  cent  alcohol  to  any  stain. 

DEHYDRATING 

After  the  sections  have  been  stained,  they  must  be  dehydrated. 
If  they  have  been  stained  in  a  strong  alcoholic  solution,  transfer  to 
95  and  then  to  100  per  cent  alcohol,  5  minutes  in  each,  if  the  stain  does 
not  wash  out  too  rapidly.  If  stained  in  an  aqueous  solution,  pass 
through  the  series,  water,  35,  50  ,70,  85,  95,  and  100  per  cent  alcohol, 
about  5  minutes  in  each. 

With  stains  which  wash  out  rapidly,  the  times  must  be  shortened 
and  some  of  the  alcohols  must  be  omitted.  With  aqueous  gentian- 
violet,  all  must  be  omitted  except  the  95  and  100  per  cent,  and  even 
in  these  the  time  must  be  shortened  to  a  few  seconds. 

CLEARING 

After  the  sections  have  been  dehydrated,  they  must  be  cleared, 
or  made  transparent  by  some  clearing  agent.  The  clearing  agent 
must  be  a  solvent  of  balsam,  but  it  is  not  at  all  necessary  that  the 
balsam  shall  be  dissolved  in  the  particular  clearing  agent  which  has 
been  used.  Xylol  balsam  is  used  not  only  when  preparations  have 
been  cleared  in  xylol,  but  also  when  they  have  been  cleared  in  clove 
oil,  cedar  oil,  bergamot  oil,  or  other  clearing  agents. 

Xylol  is  the  most  generally  useful  clearing  agent.  Place  the  slide 
in  equal  parts  of  xylol  and  absolute  alcohol  and  then  in  pure  xylol, 
allowing  each  to  act  for  about  5  minutes. 


The  Paraffin  Method  117 

Clove  oil  is  also  an  excellent  clearing  agent.  The  clove  oil  should 
follow  the  absolute  alcohol,  without  any  mixtures.  Pour  on  a  few 
drops  of  clove  oil,  and  drain  them  off  at  once  in  such  a  way  as  to  carry 
with  them  whatever  alcohol  may  still  remain.  Then  flood  the  slide 
repeatedly  with  clove  oil,  draining  the  clove  oil  back  into  the  bottle. 
If  judiciously  used,  50  c.c.  of  clove  oil  is  enough  to  clear  one  hundred 
preparations.  Sections  are  usually  cleared  in  a  few  seconds.  The 
only  objection  to  clove  oil  is  that  mounts  harden  slowly.  To  over- 
come this  difficulty,  the  slide  may  be  dipped  in  xylol  before  mounting 
in  balsam. 

For  clearing  sections  on  the  slide,  other  clearing  agents  are 
hardly  worth  mentioning.  ' 

MOUNTING  IN  BALSAM 

After  the  sections  are  cleared,  wipe  the  slide  on  the  side  which 
does  not  bear  the  sections.  Put  on  a  drop  of  Canada  balsam  and 
add  a  clean,1  thin  cover.  Before  the  cover  is  put  on,  pass  it  through 
the  flame  of  an  alcohol  lamp  to  remove  moisture,  for  it  would  be  a  pity 
indeed  to  injure  a  preparation  at  this  stage  of  the  process.  Add  a 
label,  and  the  mount  is  complete. 

A  TENTATIVE  SCHEDULE  FOR  PARAFFIN  SECTIONS 
It  will  be  useful  to  give  several  tentative  schedules  for  the  use 
of  beginners.  It  cannot  be  too  strenuously  insisted  that  these 
schedules  are  only  tentative,  their  sole  object  being  to  give  the  beginner 
a  start.  The  following  is  a  tentative  schedule  for  the  ovary  of  a  lily 
at  any  period  before  fertilization.  The  pieces  should  not  be  more 
than  12mm.  in  length. 

1.  Chromo-acetic  acid,  1  day. 

2.  Wash  in  water,  1  day. 

3.  2£,  5,  7J,  10,  15,  20,  30,  and  50  per  cent  alcohol,  4  hours  each;   70, 
85,  and  95  per  cent  alcohol,  10  hours  each;  absolute  alcohol,  12  to 
24  hours,  changing  two  or  three  times. 

»  Slides  and  covers  should  be  treated  with  hydrochloric  acid,  or  equal  parts  of  hydro- 
chloric acid  and  water,  for  several  hours.  They  should  then  be  thoroughly  rinsed  in 
water  and  wiped  with  a  cloth  perfectly  free  from  lint.  After  rinsing  in  water,  they  may  be 
kept  in  95  per  cent  alcohol.  When  a  cover  is  needed  for  use,  it  is  Dr.  Land's  practice  to 
rest  the  corner  of  the  cover  on  a  piece  of  filter  paper  to  remove  the  drop  of  alcohol;  then 
pass  the  cover  through  the  flame  of  a  Bunsen  or  alcohol  lamp.  The  film  of  alcohol  will 
burn  and  the  cover  may  warp,  but  it  will  usually  straighten,  and  it  will  be  clean  and  dry. 


118  Methods  in  Plant  Histology 

4.  Mixtures  of  absolute  alcohol  and  xylol;  2J,  5,  10,  15,  25,  50,  75,  and 
100  per  cent  xylol,  3  or  4  hours  in  each  grade.     Change  the  pure 
xylol  once  or  twice. 

5.  Paraffin  and  xylol,  48  hours. 

6.  Melted  paraffin  in  the  bath,  30  to  40  minutes. 

7.  Imbed. 

8.  Section;  about  10/i  is  a  good  thickness. 

9.  Fasten  to  the  slide. 

10.  Dissolve  off  the  paraffin  in  xylol. 

11.  Xylol  and  absolute  alcohol,  equal  parts,  5  minutes;  100,  95,  85,  and 
70  per  cent  alcohol,  5  minutes  each. 

12.  Stain  in  safranin  (alcoholic),  6  hours  or  over  night. 

13.  Rinse  in  50  per  cent  alcohol,  using  a  trace  of  HC1  if  necessary;  then 
in  70,  85,  95,  and  100  per  cent  alcohol,  5  minutes  each. 

14.  Stain  in  gentian-violet  dissolved  in  clove  oil  (or  in  clove  oil  with  a 
little  absolute  alcohol),  10  minutes. 

15.  Treat  with  pure  clove  oil  until  the  gentian-violet  stain  is  satisfactory. 

16.  Rinse  in  xylol,  1  minute. 

17.  Mount  in  balsam. 

18.  Label. 

That  the  paraffin  method  is  tedious  and  complicated  is  uni- 
versally recognized.  Many  substitutes  have  been  tried,  but  without 
enough  success  to  justify  even  a  reference. 


CHAPTER  X 
THE  CELLOJDIN  METHOD 

The  celloidin  method  is  used  more  extensively  by  zoologists  than 
by  botanists.  Where  many  mounts  are  necessary  and  only  a  single 
section  is  needed  for  each  mount,  the  method  is  to  be  recommended, 
if  the  sections  cannot  be  cut  equally  well  without  any  imbedding. 
All  the  sections  can  be  stained  and  cleared  at  one  time,  so  that,  in 
making  the  individual  mounts,  it  is  necessary  only  to  place  a  section 
on  the  slide  and  add  a  drop  of  balsam  and  a  cover.  Another  advan- 
tage, and  the  only  one  so  far  as  the  botanist  is  concerned,  is  that 
hard  roots  and  stems,  which  cannot  be  handled  by  the  paraffin 
method,  are  cut  easily  in  celloidin.  Where* serial  sections  are  neces- 
sary, as  in  most  morphological  and  cytological  work,  the  method  is 
too  tedious  to  be  worth  even  a  trial,  unless  the  sections  cannot  be  cut 
in  any  other  way.  Besides,  most  of  the  more  valuable  stains  color 
the  celloidin  matrix,  and  if  the  matrix  be  removed,  the  more  delicate 
elements  may  be  displaced  or  even  lost. 

Celloidin  and  collodion  are  forms  of  nitro-cellulose.  They  are 
inflammable,  but  do  not  explode.  Schering's  celloidin,  which  is  only 
a  collodion  prepared  by  a  patented  process,  is  in  general  use  for 
imbedding.  Granulated  and  shredded  forms  of  celloidin  are  on  the 
market,  but  the  tablets  are  more  convenient.  Directions  for  making 
the  various  solutions  accompany  the  celloidin.  To  make  a  2  per 
cent  solution,  add  to  one  tablet  enough  ether-alcohol  to  make  the 
whole  weigh  2,000  g.  To  make  a  4  per  cent  solution,  add  another 
tablet,  and  to  make  a  6  per  cent  solution,  add  an  additional  tablet, 
and  so  on. 

The  collodion  method  was  published  by  Duval1  in  1879.  Cel- 
loidin was  recommended  by  Merkel  and  Schiefferdecker2  in  1882. 
The  principal  features  of  the  method  are  as  follows:  Material  is 

1  Duval,  Journal  de  I'anatomie,  1879,  p.  185. 

'Merkel  and  Schiefferdecker,  Archivfur  Anatomie  und  Physiologie,  1882. 

119 


120  Methods  in  Plant  Histology 

dehydrated  in  absolute  alcohol;  treated  with  ether-alcohol;  infil- 
trated with  celloidin;  imbedded  in  celloidin;  hardened  in  chloro- 
form or  alcohol;  after  which  it  .is  cut,  stained,  and  mounted. 

Eycleshymer,  who  brought  the  celloidin  method  to  a  high  degree 
of  efficiency,  published  in  1892  a  short  account,  which  may  be  sum- 
marized as  follows:  Put  the  celloidin  tablet,  or  fragments,  into  a 
wide-mouthed  bottle,  and  pour  on  enough  ether-alcohol  (equal  parts 
ether  and  absolute  alcohol)  to  cover  the  celloidin.  Occasionally 
shake  and  add  a  little  more  ether-alcohol  until  the  celloidin  is  all 
dissolved.  The  process  may  require  several  days.  The  solution 
should  have  the  consistency  of  a  very  thick  oil.  Label  this  solution 
No.  4.  Solution  No.  3  is  made  by  mixing  two  parts  of  solution  "No.  4 
with  one  part  of  ether-alcohol.  Solution  No.  2  is  made  by  mixing 
two  parts  of  No.  3  with  one  part  of  ether-alcohol.  Solution  No.  1 
consists  of  equal  parts  of  ether  and  absolute  alcohol. 

After  dehydrating,  the  material  is  placed  successively  in  solutions 
1,  2,  3,  and  4.  For  an  object  2mm.  square,  24  hours  in  each  solu- 
tion is  sufficient;  for  the  brain  of  a  cat,  a  week  is  not  too  long. 

A  paper  tray  may  be  used  for  imbedding.  Pour  the  object,  with 
the  thick  solution,  into  the  tray  and  harden  in  chloroform  for  24 
hours;  then  cut  away  the  paper  and  place  the  block  in  70  per  cent 
alcohol  for  a  few  hours.  The  material  may  be  left  indefinitely  in  a 
mixture  of  equal  parts  of  95  per  cent  alcohol  and  glycerin. 

Before  cutting,  the  object  is  mounted  upon  a  block  of  wood.  A 
block,  suited  to  the  microtome  clamp,  is  dipped  in  ether-alcohol, 
which  removes  the  air  and  insures  a  firmer  mounting.  Dip  the 
block  of  wood  in  solution  No.  3,  and  the  piece  of  celloidin  containing 
the  object  in  solution  No.  1.  Press  the  two  firmly  together,  and  place 
in  chloroform  until  the  joint  becomes  hardened. 

Set  the  blade  of  the  microtome  knife  as  obliquely  as  possible. 
Both  the  object  and  the  knife  should  be  kept  flooded  with  70  per 
cent  alcohol,  and  the  sections,  as  they  are  cut,  should  be  transferred 
to  70  per  cent  alcohol. 

Stain  in  Delafield's  haematoxylin  for  5  to  30  minutes.  Wash  in 
water  for  about  5  minutes,  and  then  decolorize  in  acid  alcohol  (2  to  5 
drops  of  hydrochloric  acid  to  100  c.c.  of  70  per  cent  alcohol)  until 


The  Celloidin  Method  121 

the  stain  is  extracted  from  the  celloidin,  or  at  least  until  the  celloidin 
retains  only  a  faint  pinkish  color.  Wash  in  70  per  cent  alcohol  (not 
acid)  until  the  characteristic  purple  color  of  the  haematoxylin  replaces 
the  red  due  to  the  acid.  Stain  in  eosin  (preferably  a  1  per  cent  solu- 
tion in  70  per  cent  alcohol)  for  2  to  5  minutes.  Dehydrate  in  95  per 
cent  alcohol  for  about  5  minutes.  Absolute  alcohol  must  not  be 
used,  unless  it  is  desirable  to  remove  the  celloidin  matrix.  Eycleshy- 
mer's  clearing  fluid  (equal  parts  of  cedar  oil,  bergamot  oil,  and  car- 
bolic acid)  clears  readily  from  95  per  cent  alcohol.  Mount  in 
balsam. 

If  serial  sections  are  necessary,  arrange  the  sections  upon  a  slide, 
using  enough  70  per  cent  alcohol  to  keep  the  sections  moist,  but  not 
enough  to  allow  them  to  float.  Cover  the  sections  with  a  strip  of 
toilet  paper,  which  can  be  kept  in  place  by  winding  with  fine  thread. 
After  the  sections  have  been  stained  and  cleared,  remove  the  excess 
of  clearing  fluid  by  pressing  rather  firmly  with  a  piece  of  blotting- 
paper.  Then  remove  the  toilet  paper  and  mount  in  balsam. 

With  occasional  slight  modifications,  we  have  used  the  method 
as  presented  by  E3^cleshymer  in  his  classes.  Instead  of  the  graded 
series  of  celloidin  solutions,  we  use  a  2  per  cent  solution,  which  is 
allowed  to  concentrate  slowly  by  removing  the  cork  occasionally, 
or  by  using  a  cork  which  does  not  fit  very  tightly.  The  material 
is  imbedded  when  the  solution  reaches  the  consistency  of  a  very 
thick  oil.  If  the  material  is  to  be  cut  immediately,  we  prefer  to 
imbed  it  and  fasten  it  to  the  block  at  the  same  time.  The  blocks 
should  have  surface  enough  to  accommodate  the  objects,  and  should 
be  about  ?  inch  thick.  White  pine  makes  good  blocks;  cork  is 
much  inferior.  Place  the  block  for  a  moment  in  ether-alcohol  and 
then  dip  into  the  2  per  cent  celloidin  the  end  of  the  block  which  was 
left  rough  by  the  saw.  With  the  forceps  remove  a  piece  of  the 
material  from  the  thick  celloidin  and  place  it  upon  the  block,  taking 
care  to  keep  it  right  side  up.  Dip  the  block  with  its  object  first  in 
thick  celloidin,  then  in  thin,  and  after  exposing  to  the  air  for  a  few 
minutes  drop  it  into  chloroform,  where  it  should  remain  for  about 
10  to  20  hours.  It  should  then  be  placed  in  equal  parts  of  glycerin 
and  95  per  cent  alcohol,  where  it  may  be  kept  indefinitely.  If  the 


122  Methods  in  Plant  Histology 

material  is  hard,  like  many  woody  stems,  it  will  cut  better  after 
remaining  in  this  mixture  for  a  couple  of  weeks. 

The  following  schedules,  beginning  with  the  celloidin  sections 
in  70  per  cent  alcohol,  will  give  the  student  a  start  in  the  staining: 

Delafield's  Haematoxylin  and  Eosin.— 

1.  70  per  cent  alcohol,  2  to  5  minutes. 

2.  Delafield's  haematoxylin,  5  to  30  minutes. 

3.  Wash  in  water,  5  minutes. 

4.  Acid  alcohol  (1  c.c.  hydrochloric  acid+100  c.c.  of  70  per  cent  alcohol) 
until  the  stain  is  extracted  from  the  celloidin,  or  at  least  until  only  a 
faint  pinkish  color  remains. 

5.  Wash  in  70  per  cent  alcohol  (not  acid)  until  the  purple  color  returns. 

6.  Stain  in  eosin  (preferably  a  1  per  cent  solution  in  70  per  cent  alcohol), 
2  to  5  minutes. 

7.  Dehydrate  in  95  per  cent  alcohol,  2  to  5  minutes.     Do  not  use  abso- 
lute alcohol  unless  you  wish  to  dissolve  the  celloidin,  which  is  not 
necessary  with  this  staining. 

8.  Clear  in  Eycleshymer's  clearing  fluid,  usually  1  to  2  minutes,  but 
sometimes  5  to  10  minutes. 

9.  Mount  in  balsam. 

Safranin  and  Delafield's  Haematoxylin. — 

1.  70  per  cent  alcohol,  2  to  5  minutes. 

2.  Safranin  (alcoholic),  6  to  24  hours. 

3.  Acid  alcohol  (a  few  drops  of  hydrochloric  acid  in  70  per  cent  alcohol) 
until  the  safranin  is  removed  from  the  cellulose  walls. 

4.  Wash  in  50  per  cent  alcohol,  5  to  10  minutes  to  remove  the  acid. 

5.  Delafield's  haematoxylin,  2  to  5  minutes. 

6.  Wash  in  water,  5  minutes. 

7.  Acid  alcohol,  a  few  seconds. 

8.  Dehydrate  in  95  per  cent  alcohol,  2  to  5  minutes,  then  in  absolute 
alcohol,  2  to  5  minutes,  which  will  partially  dissolve  the  celloidin. 

9.  Clear  in  clove  oil,  which  will  complete  the  removal  of  the  celloidin. 
10.  Be  sure  that  the  sections  are  free  from  fragments  of  celloidin  and  then 

mount  in  balsam. 

Jeffrey's  improvements  in  the  celloidin  method  have  been 
described  in  considerable  detail  by  Plowman.1  Sections  of  hard 
stems  and  roots  cut  by  this  method  could  hardly  be  surpassed, 

1  Plowman,  A.  B.,  The  Celloidin  Method  with  Hard  Tissues.  Botanical  Gazette,  37: 
456-461,  1904. 


The  Celloidin  Method  123 

and   they   are   perfectly   adapted  to   the   requirements  of  photo- 
micrography.    The  following  is  a  brief  abstract  of  Plowman's  paper: 

1.  Preparation  of  Material. — Dead  and  dry  material  should  be 
repeatedly  boiled  in  water  and  cooled  to  remove  air.     An  air-pump 
may  be  used  in  addition.     Living  material  may  be  fixed  in  a  mixture 
of  picric  acid,  mercuric  chloride,  and  alcohol: 

Mercuric  chloride,  saturated  solution,  in  30  per  cent  alcohol  .      .     3  parts 
Picric  acid,  saturated  solution,  in  30  per  cent  alcohol    ....     1  part 

Fix  24  hours,  and  wash  by  passing  through  40,  50,  60,  70,  and  80 
per  cent  alcohol,  allowing  each  to  act  for  24  hours. 

2.  Desifilication,  etc. — Silica   and   other  mineral   deposits   are 
removed  by  treating  with  .a  10  per  cent  aqueous  solution  of  com- 
mercial hydrofluoric  acid.     The  material  is  transferred  to  this  solu- 
tion from  water  or  from  the  80  per  cent  alcohol.     The  process  may 
require  3  or  4  days,  with  one  or  two  changes  of  the  acid  and  frequent 
shaking  of  the  bottle.     An  ordinary  wide-mouthed  bottle,  coated 
internally  with  hard  paraffin,  should  be  prepared,  since  the  acid  is 
usually  sold  in  bottles  with  narrow  necks.     The  bottles  are  easily 
prepared  by  filling  them  with  hot  paraffin  and  simply  pouring  the 
paraffin  out.     Enough  will  stick  to  the  bottle  to  protect  the  glass 
from  the  acid.     Wash  in  running  water  3  or  4  hours. 

3.  Dehydration.— Use  30,  50,  70,  90,  and  100  per  cent  alcohol, 
allowing  12  hours  in  each  grade. 

4.  Infiltration  with  Celloidin.— There  should  be  ten  grades  of 
celloidin:   2,  4,  6,  8,  10,  12,  14,  16,  18,  and  20  per  cent.     Transfer 
from  absolute  alcohol  to  the  2  per  cent  celloidin.     (We  should  prefer 
a  previous  treatment  with  ether-alcohol.)     The  bottle  should  be 
nearly  filled,  and  the  stopper  should  be  clamped  or  wired  in  place. 
Put  the  bottle  on  its  side  in  a  paraffin  bath  at  50°  to  60°  C.  for  12  to 
18  hours.     Cool  the  bottle  quickly  in  cold  water,  taking  care  that 
the  water  does  not  get  into  the  bottle.     Pour  out  the  2  per  cent 
solution  (which,  as  well  as  all  the  other  solutions,  may  be  used  repeat- 
edly), and  replace  it  with  the  4  per  cent,  and  proceed  in  the  same  way 
with  the  other  grades.     When  the  20  per  cent  solution  is  reached,  a 
further  thickening  is  gained  by  adding  a  few  chips  of  dry  celloidin 
from  time  to  time  until  the  mixture  is  quite  stiff  and  firm.     Remove 


124  Methods  in  Plant  Histology 

each  block  with  the  celloidin  adhering  to  it  and  harden  it  in  chloro- 
form for  12  hours.  Then  transfer  to  a  mixture  of  equal  parts  of 
glycerin  and  95  per  cent  alcohol,  where  the  material  should  remain 
for  a  few  days  before  cutting. 

Cutting,  Staining,  and  Mounting. — Although  10  /j.  is  usually 
thin  enough,  sections  are  readily  cut  as  thin  as  5  ju  by  this  method. 
Remove  the  celloidin  before  staining  by  treating  10  to  15  minutes 
with  ether;  then  wash  in  95  per  cent  alcohol  and  transfer  to  water, 
and  then  to  the  stain.  Stain  to  a  fairly  dense  purple  in  an  aqueous 
solution  of  Erlich's  haematoxylin;  wash  in  dilute  aqueous  solution  of 
calcium  or  sodium  carbonate,  and  then  in  two  changes  of  distilled 
water.  Add  a  few  drops  of  alcoholic  solution  of  equal  parts  of 
Grubler's  alcoholic  and  aqueous  safranin,  and  stain  to  a  rich  red. 
A  dilute  stain  acting  1  to  2  hours  is  better  than  a  more  concentrated 
stain  acting  for  a  shorter  time.  Transfer  directly  to  absolute  alcohol , 
clear  in  xylol,  and  mount  in  balsam. 

Haidenhain's  iron-haematoxylin  is  a  very  satisfactory  stain  for 
photographic  purposes. 

The  celloidin  method  has  its  disadvantages  as  well  as  its  advan- 
tages. It  is  extremely  slow  and  tedious,  and  it  is  rarely  possible  to 
cut  sections  thinner  than  10  /*,  while,  on  the  other  hand,  it  gives 
smoother  sections. 

Succulent  tissues,  which  are  usually  damaged  by  the  paraffin 
method,  are  easily  handled  without  any  injury  in  celloidin.  The 
fact  that  the  method  may  be  used  without  heat  is  often  a  further 
advantage.  Stems  and  roots  which  cannot  be  handled  at  all  in 
paraffin  cut  well  in  celloidin,  and  much  larger  sections  can  be  cut  than 
in  paraffin,  but  most  material  of  this  kind  can  be  cut  without  any 
imbedding. 

When  material  is  to  be  imbedded,  use  celloidin  as  a  last  resort. 
Use  paraffin  when  you  can,  celloidin  when  you  must. 


CHAPTER  XI 
SPECIAL  METHODS 

While  the  methods  already  described  are  sufficient  for  most  of 
the  routine  work  of  the  ordinary  laboratory,  special  methods  are 
often  necessary  for  special  cases.  In  nearly  every  piece  of  research 
work  the  investigator  will  find  some  modification  necessary  before 
he  can  secure  the  best  results.  A  few  methods  designed  to  meet 
special  difficulties  are  given  in  this  chapter. 

VERY  LARGE  SECTIONS 

It  is  sometimes  desirable  to  cut  very  large  sections.  Sections 
as  large  as  a  cornstalk  may  be  cut  freehand  or  in  celloidin.  A  section 
of  a  stem  of  Zamia  5  or  6  cm.  in  diameter  is  difficult  to  handle  by  the 
usual  methods.  If  a  large  microtome,  such  as  is  used  inputting 
complete  sections  of  large  brains,  is  available,  the  piece  of  stem  is 
easily  held  for  the  cutting.  Some  of  the  medium-sized  sliding 
microtomes  now  have  a  rigid  clamp  which  will  grip  a  block  3  cm. 
square.  The  lower  part  of  the  piece  can  then  be  trimmed  to  fit  the 
clamp,  leaving  the  upper  part  round,  so  that  sections  across  a  stem 
6  or  7  cm.  in  diameter  may  be  cut  without  much  difficulty.  With  a 
rather  soft  stem,  like  Zamia,  the  surface  must  be  flooded  with  95  per 
cent  alcohol  after  each  section,  if  it  is  desirable  to  cut  thin  sections. 
From  stems  3  cm.  in  diameter,  sections  can  be  cut  at  about  20  to 
30  /z.  If  the  section  is  not  more  than  3  or  4  cm.  in  diameter,  it  can 
be  mounted  on  a  50X75  mm.  slide.  Sections  6  or  7  cm.  in  diameter 
can  be  mounted  on  lantern  slides;  if  large  covers  are  not  available, 
use  another  lantern  slide  for  a  cover.  It  will  be  easier  to  get  neat 
mounts  if  the  cover  is  cut  down  so  as  to  leave  a  margin  2  or  3  mm. 
wide.  It  is  not  easy  to  mount  a  thick  section  between  two  lantern 
slides  of  the  same  size,  on  account  of  the  balsam  which  oozes  out  at 
the  edges.  Such  preparations  may  be  used  directly  as  lantern  slides. 
Large  sections  of  the  stem  of  a  tree  fern  make  good  mounts  without 

125 


126  Methods  in  Plant  Histology. 

any  staining.  Large  sections  of  cornstalk  are  rather  hard  to  cut, 
because  the  rigid  bundles  tear  through  the  soft  parenchyma.  Flood- 
ing with  95  per  cent  alcohol  facilitates  the  process.  A  slight  harden- 
ing is  sufficient,  So  that  about  4  or  5  sections  can  be  cut  in  1  minute. 

STONY  TISSUES 

Sections  of  the  stony  tissues  of  hickory  nuts,  walnuts,  peach 
stones,  and  similar  refractory  substances  cannot  be  cut  by  ordinary 
methods. 

With  a  fine  saw,  saw  sections  about  1  mm.  in  thickness.  Rub 
a  section  between  two  pieces  of  fine  sandpaper  until  it  is  not  more 
than  half  a  millimeter  in  thickness.  Then  rub  it  between  two  smooth 
hones,  keeping  the  hones  wet  with  water.  When  the  section  is  thin 
enough,  wash  it  thoroughly  in  water,  using  a  pipette  to  rinse  off  any 
particles  of  dirt.  Dehydrate  in  absolute  alcohol,  clear  in  clove  oil, 
and  mount  in  balsam.  The  long,  narrow  pores  show  better  without 
any  clearing.  In  this  case,  dry  the  section  thoroughly,  heat  a  few 
drops  of  balsam  on  the  slide  to  drive  off  the  solvent,  put  the  section 
into  the  balsam,  and  add  a  cover.  The  air  caught  in  the  long,  narrow' 
pores  will  make  them  appear  as  black  lines.  Sections  of  most  nuts 
show  excellent  detail  without  any  staining.  Thin  sections,  however, 
may  be  stained  in  the  usual  way. 

PETRIFACTIONS 

During  the  past  ten  years  the  study  of  fossil  plants  has  made 
even  more  rapid  progress  than  during  the  previous  decade.  Scarcely 
any  problem  involving  the  anatomy  of  living  vascular  plants  can  be 
investigated  intelligently  without  some  knowledge  of  Paleozoic  and 
Mesozoic  forms.  Consequently,  it  is  becoming  increasingly  neces- 
sary for  laboratories  to  have  apparatus  and  technic  for  cutting  rock- 
sections. 

The  outline  of  the  process  of  cutting  a  rock-section  is  very  simple : 

1.  Saw  the  rock  into  two  piece?, 

2.  Polish  the  cut  surface. 

3.  Fasten  the  cut  surface  to  a  piece  of  glass  with  hot  shellac. 

4.  With  the  saw,  make  another  cut,  as  close  to  the  glass  as  possible, 
so  as  to  leave  a  thin  section  firmly  fastened  to  the  glass. 


Special  Methods  127 

5.  Grind  and  polish  until  the  section  is  as  thin  as  possible,  or  as  thin  as 
you  want  it. 

6.  Wash  all  polishing  powder  off  with  water. 

7.  Dry  completely  and,  either  with  or  without  moistening  in  xylol,  mount 
in  balsam. 

A  word  of  suggestion  in  regard  to  these  various  points  may  not 
be  amiss. 

1.  Most   rock  sections   are   cut  with   a  rather  expensive  and 
quite  complicated  instrument,   called   a  petrotome.    The  saw  is 
of   the  circular   type,   is  made  of  tin   or  other  soft  metal,   has 
no  teeth,  but  has  diamond  dust  driven  into  the  margin.     A  rigid 
clamp  holds  the  object,  and  the  saw,  driven  at  a  great  speed  and 
constantly  cooled  by  a  stream  of  water,  gradually  cuts  through 
the  specimen. 

2.  The  cut  surface  is  most  easily  polished  on  a  revolving  brass 
plate,   kept  wet  and  liberally  powdered  with  fine  carborundum. 
When  the  surface  has  become  even  and  smooth,  the  specimen  is 
ready  for  the  next  step. 

3.  Fasten  to  the  glass  slide  upon  which  the  section  js  to  be 
mounted.     Plate  glass  3  or  4  mm.  thick  is  best  for  sections  larger 
than  3  or  4  mm.  square.     Gradually  heat  the  slide  until  it  is  quite 
hot.     Melt  upon  the  slide  the  thin  brown  or  white  shellac  used  by 
painters;   heat  the  object  and  press  the  polished  surface  very  firmly 
into  the  melted  shellac.     As  soon  as  the  slide  and  object  are  cool,  the 
next  cut  can  be  made. 

4.  Anyone  who  can  handle  tools  should  soon  be  able  to  cut  a 
section  1  mm.  thick.     A  skilled  technician  can  cut  sections*  as  thin 
as  0.5  mm. 

5.  The  second  grinding  must  be  very  careful  and  accurate.     Do 
the  polishing  on  the  revolving  disk.     The  glass  slide  allows  one  to 
note  how  the  process  is  progressing. 

6.  When  the  section  becomes  thin  enough,  or  even  before  if  it 
begins  to  crack,  wash  off  the  powder.     If  the  slide  has  been  damaged 
and  the  section  is  holding  together,  the  shellac  may  be  dissolved 
with  absolute  alcohol,  thus  freeing  the  section,  which  may  now  be 
mounted  on  another  slide. 


128  Methods  in  Plant  Histology 

7.  It  is  usually  a  good  plan  to  use  rather  thick  balsam  for  mount- 
ing, even  if  it  should  be  necessary  to  heat  it  a  little  to  make  it  flow 

well. 

By  this  method,  sections  of  silicious  fossils  10X15  cm.  have  been 
cut  thin  enough  for  examination  with  a  4  mm.  objective.  Sections 
3  or  4  mm.  square  have  been  cut  thin  enough  for  satisfactory  examina- 
tion with  a  2  mm.  oil  immersion  lens. 

Of  course,  this  method  can  be  used  for  such  objects  as  walnut 
and  hickory  shells. 

THICK  SECTIONS 

It  is  sometimes  desirable  to  make  very  thick  sections  to  show 
general  topography  rather  than  detail.  A  longitudinal  section  of 
the  fully  grown  ovule  of  Ginkgo  or  a  cycad  may  be  cut  as  thick  as 
3  to  5  mm.  so  as  to  include  the  entire  group  of  archegonia.  A  slab 
can  be  cut  from  each  side  of  the  ovule  with  a  fine  saw,  and  a  razor 
can  be  used  for  smoothing.  If  the  section  is  from  fresh  material  it 
should  be  fixed,  washed,  etc.,  with  about  the  same  periods  as  if  it 
were  to  be  imbedded  in  paraffin.  When  thoroughly  cleared  in  xylol, 
the  section  should  be  put  into  a  flat  museum  jar  of  suitable  size  and 
kept  in  xylol.  Even  before  the  stony  coat  of  a  cycad  becomes  too 
hard  to  be  cut  with  a  razor  such  thick  sections  are  very  instructive. 
Stain  very  lightly,  or  not  at  all.  Sections  .of  Zamia  or  other  cycad 
stems,  about  2  mm.  thick,  make  instructive  mounts,  since  they  show 
the  peculiar  course  of  the  bundles,  a  feature  which  is  largely  lost  in 
thin  sections. 

LAND'S  GELATIN  METHOD 

It  is  sometimes  desirable  to  get  sections  of  partly  disorganized 
material.  A  matrix  is  necessary  to  hold  the  parts  in  place,  but 
dehydration  may  make  the  tissue  unnecessarily  hard  to  cut. 

Soak  ordinary  gelatin  (which  can  be  obtained  at  the  grocery)  in 
water  until  no  more  is  taken  up.  Then  drain  off  the  excess  water 
and  liquefy  the  gelatin  by  heating.  Place  the  material — previously 
soaked  in  water — in  the  melted  gelatin  and  keep  it  there  for  several 
hours.  Place  also  in  the  gelatin  some  small  blocks  of  hard  wood 
to  serve  as  supports  in  the  microtome.  The  material  to  be  sectioned 


Special  Methods  129 

is  oriented  in  a  gelatin  matrix  on  the  supporting  blocks,  cooled  until 
the  gelatin  sets,  and  then  placed  in  strong  formalin  to  harden  the 
gelatin.  In  cutting,  flood  the  knife  with  water. 

If  the  material  is  to  be  stained,  stain  it  in  bulk  before  putting  it 
into  the  gelatin,  since  the  gelatin  stains  very  deeply.  Of  course, 
the  gelatin  could  be  dissolved  with  hot  water,  or  hot  water  and  acetic 
acid,  but  all  the  advantage  of  a  matrix  would  be  lost. 

It  would  be  worth  while  to  try  this  method  thoroughly  with  soft, 
succulent  tissues  and  with  hard  tissues  which  become  still  harder 
if  dehydrated. 

SCHULTZE'S  MACERATION  METHOD 

Various  solutions  are  used  to  separate  a  tissue  into  its  individual 
cells.  These  solutions  dissolve  or  weaken  the  middle  lamella  so  that 
the  cells  are  easily  shaken  or  teased  apart.  Schultze  used  strong  nitric 
acid  and  potassium  chlorate.  Put  the  material,  which  should  be  in 
very  small  pieces,  into  a  test-tube;  pour  on  just  enough  nitric  acid 
to  cover  it,  and  then  add  a  few  crystals  of  potassium  chlorate.  Heat 
gently  until  bubbles  are  evolved,  and  let  the  reagent  act  until  the 
material  becomes  white.  Four  or  five  minutes  should  be  sufficient. 
The  fumes  are  disagreeable  and  are  very  injurious  to  microscopes. 
Pour  the  contents  of  the  tube  into  a  dish  of  water.  After  the  material 
is  thoroughly  washed  in  water,  it  may  be  teased  with  needles  and 
mounted,  or  it  may  be  put  into  a  bottle  of  water  and  shaken  until 
many  of  the  cells  become  dissociated. 

After  a  thorough  washing  in  water,  the  material  may  be  stained. 
The  large  tracheids  of  ferns,  dissociated  in  this  way  and  stained  in 
safranin  or  methyl  green,  make  beautiful  preparations. 

PROTOPLASMIC  CONNECTIONS 

In  exceptional  cases,  like  the  sieve  plates  of  the  Cucurbitaceae, 
the  protoplasmic  connections  show  plainly  with  ordinary  methods, 
but  in  most  cases  it  is  necessary  to  resort  to  special  methods  in  order 
to  demonstrate  protoplasmic  continuity.  In  these  special  methods  a 
reagent  is  used  which  causes  the  membranes  to  swell  before  the  stain 
is  applied.  It  is  only  by  such  an  exaggeration  that  the  more  delicate 
connections  can  be  shown. 


130  Methods  in  Plant  Histology 

Put  thin  sections  of  fresh  material  into  a  mixture  of  equal  parts 
of  sulphuric  acid  and  water;  and  allow  the  reagent  to  act  for  2  to 
10  seconds.  Wash  the  acid  out  thoroughly  in  water  and  stain  in 
anilin  blue.  According  to  Gardiner,  this  stain  should  be  made  by 
adding  1  g.  of  the  dry  stain  to  100  c.c.  of  a  saturated  solution  of 
picric  acid  in  50  per  cent  alcohol.  The  staining  solution  is  then 
washed  out  in  water,  and  the  sections  are  mounted  in  glycerin.  The 
sections  may  be  dehydrated,  cleared  in  clove  oil,  and  mounted  in 
balsam.  The  anilin  blue  may  be  used  in  50  per  cent  alcohol  acidu- 
lated with  a  few  drops  of  acetic  acid. 

Chloroi'odide  of  zinc  may  be  used  instead  of  sulphuric  acid.  Treat 
the  fresh  sections  for  2  hours  with  the  iodine  and  potassium-iodide 
solution  used  in  testing  for  starch;  then  treat  about  12  hours  with 
chloro'iodide  of  zinc.  Wash  in  water  and  stain  in  anilin  blue. 
Examine  in  glycerin. 

Meyer's  pyoktanin  method  is  one  of  the  best.  The  reagents  are 
as  follows: 

1.  Iodine,  potassium  iodide  solution:   iodine  1  part,  potassium  iodide 
1  part,  water  200  parts. 

2.  Sulphuric  acid  1  part,  water  3  parts;  this  mixture  to  be  saturated 
with  iodine. 

3.  Pyoktanin  coeruleum  1  g.,  water  30  c.c.    This  pyoktanin  is  a  very 
pure  methyl  violet  obtained  from  E.  Merck  in  Darmstadt. 

Put  sections  of  the  date  seed  into  a  watch  glass  full  of  the  first 
solution,  and  allow  it  to  act  for  a  few  minutes;  then  mount  in  a  drop 
of  the  solution.  The  connections  will  be  only  very  faintly  stained, 
showing  a  slightly  yellowish  color.  At  the  edge  of  the  cover,  add  a 
drop  of  the  second  solution.  The  preparation  will  darken  a  little. 
Then  allow  a  small  drop  of  the  third  solution  to  run  under  the  cover. 
Allow  the  stain  to  act  for  about  3  minutes.  Then  plunge  the 
whole  preparation  into  water.  The  action  should  be  stopped 
before  the  entire  section  has  become  blue.  Now  wash  the  section 
quickly.  If  there  are  annoying,  granular  precipitates,  remove 
them  with  a  soft  brush.  Mount  in  glycerin.  The  membrane 
should  be  a  clear  blue,  while  the  protoplast  and  connections  should 
be  a  blue  black. 


Special  Methods  131 

The  following  is  Strasburger's  modification  of  Meyer's  method, 
and  shows  the  connections  with  great  distinctness: 

1.  Treat  the  fresh  sections  with  1  per  cent  osmic  acid,  5  to  7  minutes. 

2.  Wash  in  water  5  to  10  minutes. 

3.  Treat  with  a  solution  of  iodine  in  potassium  iodide  (0.2  per  cent 
iodine  and  1.64  per  cent  potassium  iodide),  20  to  30  minutes. 

4.  Transfer  to  25  per  cent  sulphuric  acid,  which  should  act  for  at  least 
half  an  hour;  24  hours  may  be  necessary. 

5.  Bring  the  sections  into  25  per  cent  sulphuric  acid  which  has  been 
saturated  with  iodine.     Add  a  drop  of  Meyer's  pyoktanin  solution 
(1  g.  pyoktanin  coeruleum  as  sold  by  E.  Merck  in  Darmstadt  in  30  c.c. 
of  water). 

In  about  5  minutes  the  sections  will  be  stained  sufficiently  and 
can  be  examined  in  glycerin.  If  there  are  annoying  precipitates, 
remove  them  with  a  soft  brush. 

According  to  Meyer,  the  swelling  is  an  advantage  only  when 
the  walls  are  very  thin.  When  the  walls  are  thick,  the  connections 
show  better  without  any  previous  swelling. 

Try  the  following  method  with  the  seeds  of  Diospiros,  Latania, 
Chamerops,  Phoenix,  or  Phytelephas:  Soak  in  water  and  cut  thin 
sections.  Extract  the  oily  and  fatty  substances  with  xylol;  wash 
in  95  per  cent,  or  in  absolute  alcohol;  stain  in  anilin  blue  (Hoffman's 
blue  1  g.  dissolved  in  150  c.c.  of  50  per  cent  alcohol)  for  a  few  minutes. 
Examine  in  glycerin.  This  method  succeeds  very  well  with  seeds  of 
the  date,  which  is  sold  at  all  groceries. 

Permanent  preparations  may  be  secured  by  the  following  method : 

1.  Fix  in  1  per  cent  osmic  acid,  or  in  absolute  alcohol,  5  to  10  minutes. 

2.  Stain  for  24  hours  in  Delafield's  haematoxyiin. 

3.  Wash  for  a  few  minutes  in  acid  alcohol  (5  drops  of  hydrochloric  acid 
in  50  c.c.  of  70  per  cent  alcohol). 

4.  Wash  for  a  few  minutes  in  ammonia  alcohol  (5  drops  of  ammonia  to 
50  c.c.  of  70  per  cent  alcohol). 

5.  Dehydrate  in  absolute  alcohol,  clear  in  xylol,  and  mount  in  balsam. 

STAINING  CILIA 

The  cilia  of  the  large  spermatogoid  of  Ginkgo  and  the  cycads 
take  a  brilliant  stain  with  gentian-violet,  whether  the  gentian-violet 
be  used  alone  or  in  combination  with  safranin.  The  cilia  of  the 


132  Methods  in  Plant  Histology 

spermatozoids  of  the  pteridophytes  also  stain  by  this  method, 
although  not  so  brilliantly  as  in  case  of  the  cycads. 

The  cilia  of  the  motile  spores  of  Thallophytes  may  often  be 
demonstrated  by  allowing  a  drop  of  the  iodine  solution  used  in  testing 
for  starch  to  run  under  the  cover. 

Zimmermann  gives  the  following  method:  Bring  the  objects  into 
a  drop  of  water  on  the  slide  and  invert  the  drop  over  the  fumes  of  1 
per  cent  osmic  acid  for  5  minutes.  Allow  the  drop  to  dry.  Then  add 
a  drop  of  20  per  cent  aqueous  solution  of  tannin,  and  after  5  minutes 
wash  it  off  with  water.  Stain  in  a  strong  aqueous  solution  of  f uchsin 
(or  carbol  fuchsin)  for  5  minutes.  Allow  the  preparation  to  dry 
completely,  and  then  add  a  drop  of  balsam  and  a  cover.  The  cilia 
should  take  a  bright  red. 

Zimmermann  also  found  the  following  method  satisfactory  for 
the  cilia  of  the  zoospores  of  algae  and  fungi:  Fix  by  adding  a  few 
drops  of  1  per  cent  osmic  acid  to  the  water  containing  the  zoospores; 
then  add  an  equal  amount  of  a  mixture  of  fuchsin  and  methyl  violet. 
The  fuchsin  and  methyl  violet  should  be  1  per  cent  solutions  in  95  per 
cent  alcohol.  In  a  few  seconds  the  cilia  stain  a  bright  red. 

While  gentian-violet  gives  the  cilia  of  cycads  a  beautiful  and 
brilliant  stain,  we  have  found  that  nothing  surpasses  Haidenhain's 
iron-alum  haematoxylin  in  giving  clear  and  definite  views  of  cilia. 

MITOCHONDRIA 

Since  the  second  edition  of  this  book  appeared,  in  1908,  the  terms 
mitochondria,  chondriosomes,  Chondriokonten,  Chondromiten,  etc., 
have  become  increasingly  frequent  in  botanical  literature.  These 
mitochondria,  as  we  shall  call  them,  are  minute  structures,  probably 
present  in  most  cells,  but  not  differentiated  by  the  most  usual 
methods  and  generally  overlooked  when  they  might  be  seen.  Most 
of  them  are  as  small  as  bacteria  and  bear  a  superficial  resemblance 
to  coccus,  spirillum,  and  bacillus  forms  (Fig.  24,  A). 

Many  fixing  agents  either  destroy  the  mitochondria  or  make  it 
almost  impossible  to  demonstrate  them.  Fixing  agents  containing 
alcohol  or  any  considerable  percentage  of  acid  are  to  be  avoided. 
Benda's  solution,  followed  by  Haidenhain's  iron-alum  haematoxylin, 


Special  Methods 


133 


will  give  good  results.     A  solution  recommended  by  Bensley  is  good 
also  for  plant  material. 

Bensley's  Solution. — 

Osmic  acid  2  per  cent 1  part 

Corrosive  sublimate  (HgCl2)  2£  per  cent 4  parts 

Add  one  drop  of  glacial  acetic  acid  to  10  c.c.  of  this  solution. 
Fix  for  24  to  48  hours  and  wash  thoroughly  in  water.  On  the  slide, 
bleach  with  hydrogen  peroxide;  wash  in  water;  treat  with  the  iodine 


FIG.  24.— Cells  from  the  periblem  of  the  root-tip  of  Allium  cepa:  A,  mitochrondria; 
B,  canaliculi ;  fixed  in  Bensley's  solutions  and  stained  in  iron-alum  haematoxylin.  X 1200. 

solution  used  in  testing  for  starch;  then  wash  in  water.  The  slide 
is  now  ready  for  staining.  We  recommend  the  usual  Haidenhain's 
iron-alum  haematoxylin. 

Bensley  recommends  the  following  method  which  we  have  found 
rather  uncertain,  but  which,  when  successful,  yields  magnificent 
preparations:  On  the  slide,  bleach  for  2  or  3  seconds  in  a  1  per  cent 
aqueous  solution  of  permanganate  of  potash;  then  treat  with  a 
5  per  cent  aqueous  solution  of  oxalic  acid  until  the  preparation 
becomes  white  (a  few  seconds);  wash  in  water,  and  then  stain  as 
follows : 

1.  Copper  acetate  (neutral)  saturated  solution  in  water,  5  to  10  minutes. 

2.  Wash  in  water. 

3.  %  per  cent  haematoxylin,  5  to  10  minutes. 

4.  Wash  in  water. 

5.  Potassium  bichromate  (neutral)  5  per  cent  solution  in  water  until  the 
preparation  blackens,  usually  30  seconds  or  less. 


134  Methods  in  Plant  Histology 

6.  Differentiate  in  Weigert's  ferricyanide  solution. 

Borax •••       2-Og- 

Ferricyanide  of  potassium 2.5  g. 

Water 200.0c.c. 

7.  Wash  in  water  and  proceed  as  usual. 

CANALICULI 

By  using  special  methods,  Bensley  has  obtained  views  of  the 
protoplasm  of  plants,  quite  different  from  those  seen  in  ordinary 
preparations.  In  the  cell  of  a  root-tip  a  series  of  small  canals,  or 
vacuoles,  appears,  which  is  much  more  definite  and  extensive  than 
the  usual  display  of  vacuoles  and  which  appears  before  any  vacuoles 
can  be  recognized  in  preparations  made  in  the  usual  way  (Fig.  24,  B). 
Being  a  zoologist,  he  has  called  these  vacuoles  canaliculi. 

Bensley's  Fixing  Agent. — 

1.  Formalin  (neutral) 10. 0  c.c. 

2.  Bichromate  of  potash 2.5  g. 

3.  Corrosive  sublimate 5. 0  g. 

4.  Water 90. 0  c.c. 

Dissolve  the  bichromate  of  potash  in  the  water,  then  add  the 
corrosive  sublimate  and  finally  add  the  formalin.  The  solution  2, 
3,  4  will  keep,  but  the  formalin  soon  becomes  acid.  Add  the  formalin 
to  2,  3,  4  only  when  needed  for  use.  Obtain  the  neutral  formalin 
by  distilling  the  ordinary  commercial  formalin.  Proceed  as  follows: 

1.  Fix  24  hours. 

2.  Wash  in  water,  24  hours. 

3.  Iodize  on  the  slide. 

4.  Wash  in  water,  5  minutes. 

5.  Copper  acetate  (neutral)  saturated  solution  in  water,  5  to  10  minutes. 

6.  Wash  in  water,  1  minute. 

7.  ^  per  cent  haematoxylin,  5  to  10  minutes. 

8.  Wash  in  water,  1  minute. 

9.  Potassium  bichromate  (neutral)  5  per  cent  in  water  till  it  blackens, 
about  30  seconds  or  less. 

10.  Weigert's  ferricyanide  solution  until  the  preparation  looks  right. 

11.  Wash  in  water  and  proceed  as  usual. 


Special  Methods  135 

VASCULAR  BUNDLES  IN  LIVING  TISSUES 

In  studying  venation,  and  in  tracing  the  course  of  vascular 
bundles  in  large  ovules  and  in  other  organs,  it  is  often  an  advantage 
to  use  a' stain.  If  a  stem  of  Impatiens  be  cut  under  water,  and  the 
cut  surface  be  then  placed  in  a  dilute  aqueous  solution  of  eosin,  the 
eosin  will  rise  in  the  vessels,  making  them  very  prominent.  The 
outer  bundles  of  the  large  ovules  of  cycads  are  very  easily  studied 
by  this  method.  The  inner  bundles  also  may  be  seen  by  opening 
the  seed  and  removing  the  endosperm. 

If  such  preparations  could  only  be  cleared,  they  would  be  still 
more  valuable,  but  the  effect  is  due  to  the  presence  of  the  staining 
fluid  in  the  vessels,  and  any. subsequent  treatment  diffuses  or  destroys 
the  stain.  Perhaps  a  little  experimenting  might  obviate  the  diffi- 
culty. 

STAINING  LIVING  STRUCTURES 

Some  stains  will  stain  living  structures.  Cyanin,  methyl  blue, 
and  Bismarck  brown  have  been  recommended  for  this  purpose.  The 
solutions  should  be  very  dilute,  not  stronger  than  1 : 10,000  or 
1:500,000.  The  solutions  should  be  very  slightly  alkaline,  never 
acid.  It  is  claimed  that  such  solutions  never  stain  the  nucleus,  and 
that  if  the  nucleus  stains  at  all,  it  is  an  indication  that  death  is  taking 
place. 

Campbell  succeeded  in  staining  the  living  nuclei  in  the  stamen 
hairs  of  Tradescantia  by  using  dilute  solutions  of  dahlia  and  of 
methyl  violet  (0.001  to  0.002  per  cent  in  water).  Dividing  nuclei 
were  stained. 

For  determining  the  stage  of  development  of  fresh  material  it  is 
often  necessary  to  use  a  stain.  For  this  purpose  stronger  stains  may 
be  used,  since  it  is  unimportant  whether  the  tissue  is  killed  or  not. 
An  aqueous  solution  of  methyl  green  or  eosin  can  be  recommended. 
With  1  per  cent  solutions,  diluted  one-half  with  water,  mitotic 
figures  can  be  recognized  with  ease. 


CHAPTER  XII 
PHOTOMICROGRAPHS  AND  LANTERN  SLIDES 

While  a  work  like  the  present  book  is  hardly  the  place  for  any 
extended  treatment  of  photomicrography  or  the  making  of  lantern 
slides,  a  few  simple  directions  will  help  the  beginner  and  enable  him 
to  prepare  most  of  the  photomicrographs  and  lantern  slides  which 
may  be  necessary  in  the  classroom.  It  is  assumed  that  the  student 
knows  how  to  handle  an  ordinary  camera  and  knows  how  to  do  his 
own  developing. 

PHOTOMICROGRAPHS 

For  a  simple  beginning,  no  apparatus  is  needed  except  an  ordi- 
nary camera  and  a  microscope.  Try  low  powers  first  and  proceed 
gradually  to  the  higher  magnifications.  Remove  both  front  and 
back  lenses  from  the  camera,  leaving  the  lens  barrel  and  the  shutter; 
also,  remove  the  eyepiece  from  the  microscope.  Bend  the  micro- 
scope to  the  horizontal  position  and  place  the  lens  of  the  camera 
close  to  the  ocular  end  of  the  microscope  and  shut  out  all  light  at 
this  point  by  winding  black  cloth  around  the  end  of  the  microscope 
and  the  barrel  of  the  camera  lens.  Take  great  care  to  have  a  per- 
fectly straight  optical  axis  through  the  microscope  and  camera. 

While  the  camera  and  microscope  can  be  adjusted  so  as  to  secure 
a  perfect  optical  axis  by  simply  putting  both  instruments  on  the  table 
and  raising  one  or  the  other — according  to  the  size  of  the  camera — 
by  placing  a  board  under  it,  such  an  adjustment  is  extremely  un- 
satisfactory, since  the  least  jar  may  disturb  it,  and  inserting  the 
plateholder  is  almost  sure  to  disarrange  something.  It  will  save  time 
if  you  prepare  a  board  to  keep  both  instruments  in  position.  Select 
a  clear  board  1  inch  thick,  about  1  foot  wide,  and  5  feet  long.  On 
the  top  of  this  board,  screw  two  pieces  f  inch  thick,  1|  inches  wide, 
and  5  feet  long,  so  as  to  form  a  guideway  for  the  camera  (Fig.  25,  A). 
If  the  camera  is  so  small  that  it  must  be  raised  to  bring  it  into  the 

136 


Photomicrographs  and  Lantern  Slides 


137 


optical  axis  of  the  miscroscope,  fit  to  the  guideway  a  board  of  the 
necessary  thickness,  and  fasten  the  camera  to  this  board.  It  is 
absolutely  necessary  that  the  preparation  to  be  photographed  and 
the  ground  glass  of  the  camera  should  be  perfectly  parallel.  The 
board  will  save  time  in  securing  this  parallelism.  Cut  through  the 
board  a  slot  \  inch  wide  and  extending  to  within  6  inches  of  each  eno. 
By  this  means  the  camera  can 
be  clamped  with  the  screw 
used  to  fasten  it  to  a  tripod. 
Also,  a  piece  of  metal  or  hard 
wood  may  be  placed  over  the 
horseshoe  base  of  the  micro- 
scope and  with  a  bolt,  pref- 
erably one  with  a  butterfly 
nut,  the  microscope  may  be 
held  firmly  in  place.  This 
board,  with  the  long  slot,  will 
be  useful  in  making  lantern 
slides. 

As  an  illuminant,  direct 
sunlight,  diffuse  daylight,  a 
gas-mantle  lamp,  an  acetylene 
lamp,  a  Nernst  lamp,  an  arc 
light,  or  any  strong  light  may 
be  used.  Remove  the  mirror 
from  the  microscope  and  allow 

the  light  to  come  directly  into  the  optical  axis.     This  mirror  will 
not  be  needed  in  any  photomicrographic  work. 

Let  us  suppose  that  we  are  to  make  a  photomicrograph  of  a 
vascular  bundle  and  that  we  are  using  a  16  mm.  objective.  If 
only  a  part  of  the  bundle  is  shown  on  the  ground  glass,  remove 
the  ocular  of  the  miscroscope.  If  the  illumination  is  very  uneven 
and  shows  a  "  flare  spot,"  look  at  the  inside  of  the  tube  of  the  micro- 
scope. Probably,  it  was  not  blackened  and  the  "flare  spot"  was 
due  to  reflections.  Obviate  the  difficulty  by  putting  a  piece  of 
black  paper  inside  the  tube.  Any  modern  microscope  should  have 


FIG.  25. — A,  board  for  photomicrographic 
and  lantern-slide  work;  B,  end  view  with 
clips  to  hold  negative;  C,  side  view  of  block  to 
be  used  on  board  when  making  lantern  slides. 


138  Methods  in  Plant  Histology 

the  tube  well  blackened  inside.  Move  the  light  back  and  forth  and 
sidewise  to  get  the  best  illumination.  About  six  inches  from  the 
stage  is  likely  to  be  somewhere  near  the  proper  position.  If  the 
illumination  is  still  uneven,  remove  the  condenser  from  the  micro- 
scope. The  ordinary  form  of  Abbe  condenser  is  not  likely  to  be 
satisfactory  with  objectives  of  16  mm.  focus  and  should  not  be  used 
at  all  with  objectives  of  such  long  focus. 

Focus  the  object  upon  the  ground  glass.  Even  with  a  16  mm. 
objective,  the  ordinary  ground  glass  is  rather  coarse  for  accurate 
focusing.  Always  examine  the  image  with  a  focusing  lens.  A 
brilliant  view  may  be  obtained  by  fastening  a  thin  cover-glass  to  the 
ground  glass  with  a  small  drop  of  balsam.  At  this  spot  the  image 
may  be  examined  very  critically.  Of  course,  as  in  any  photography, 
the  ground  side  of  the  glass  should  be  nearest  the  object,  occupying 
exactly  the  place  which  is  to  be  occupied  by  the  emulsion  side  of  the 
plate.  Do  not  focus  indiscriminately,  but  be  sure  that  the  image 
is  sharp  at  the  level  of  the  ground  side  of  the  glass.  It  is  a  good  plan 
to  make  a  cross  upon  the  ground  glass  with  a  pencil  or  pen,  and  then 
add  a  drop  of  balsam  and  a  cover-glass.  Focus  on  this  mark  and 
fix  the  focusing  glass  at  this  level.  The  cheap  tripod  lenses  are  good 
for  this  purpose. 

The  time  of  exposure  will  vary  with  the  magnification,  the 
intensity  of  the  light,  and  the  speed  of  the  plate.  The  exposures  will 
be  much  longer  than  in  ordinary  photography.  It  is  better  to  use 
artificial  light,  since  one  can  more  quickly  learn  to  estimate  the 
length  of  exposure  when  the  intensity  of  the  light  is  constant.  A 
slow  plate,  even  the  very  slow  contrast  plate,  is  likely  to  prove  most 
satisfactory.  With  a  Welsbach  lamp,  a  contrast  plate  of  the  same 
speed  as  a  lantern  slide,  and  a  16  mm.  objective  used  without  an 
ocular,  or  Abbe  condenser,  try  an  exposure  of  30  seconds.  Develop 
the  negative  in  whatever  solution  is  recommended  in  the  directions 
which  come  with  every  box  of  plates.  If  the  negative  is  too  weak, 
make  a  longer  exposure;  if  too  dense,  shorten  the  exposure.  A 
little  experience  with  your  apparatus  will  soon  enable  you  to  estimate 
the  length  of  exposure  with  some  certainty.  We  use  lantern  slides 
for  all  tests  and  for  small  photomicrographs.  The  Cramer  lantern 


Photomicrographs  and  Lantern  Slides  139 

slides  and  contrast  plates  of  larger  sizes  have  the  same  speed  and, 
consequently,  one  can  determine  the  length  of  exposure  by  using  a 
cheap  lantern  slide.  In  making  tests,  it  will  save  both  time  and 
money  to  expose  for  5  seconds  and  then  push  in  the  dark  slide  so  as  to 
cover  a  part  of  the  plate;  then  expose  5  seconds  longer  and  push  the 
slide  in  a  little  farther,  etc.  In  this  way  you  can  make  four  or  five 
exposures  on  a  lantern  slide  plate,  showing  exposures  of  5,  10,  15,  20, 
and  25  seconds,  the  first  exposure  being  5  seconds  and  the  last,  25 
seconds.  A  print  from  such  a  negative  is  valuable,  since  it  enables 
one  to  judge  very  accurately  the  printing  quality  of  the  various 
exposures. 

The  ordinary  filters  with  a  bichromate  of  potash  color,  used  in 
out-of-door  work,  are  good,  especially  when  used  in  addition  to  some 
filter  suited  to  the  particular  stain.  We  have  found  a  yellowish-green 
filter  very  good  for  most  iron-alum  haematoxylin  stains  and  also 
for  the  safranin,  gentian-violet,  orange  combination.  We  prefer  a 
stained-glass  filter,  because  it  is  constant  and  careful  records  will  soon 
enable  one  to  guess  with  considerable  precision,  while  liquid  filters 
vary  so  much  that  records  have  comparatively  little  value.  Filters, 
of  course,  lengthen  the  exposure.  The  strong  photographic  filter 
mentioned  in  the  descriptions  of  several  of  the  photomicrographs  in 
this  book  increases  the  exposure  15  times. 

With  fast  plates  and  without  filters,  a  strong  light  will  allow 
exposures  of  a  fraction  of  a  second,  but  we  have  had  no  success  under 
such  conditions. 

The  Abbe  condenser,  which  should  not  be  used  at  all  with  low 
powers,  is  very  useful  with  objectives  of  8  mm.  focus  and  all  higher 
powers,  especially  if  the  condenser  is  achromatic.  If  the  con- 
denser is  not  achromatic,  it  is  sometimes  a  good  plan  to  remove  it  and 
in  its  place  put  a  16  mm.  objective,  or,  for  very  high  powers,  even 
an  8  mm.  objective.  The  condenser  may  be  fastened  into  the  con- 
denser sleeve  by  an  improvised  ring  or  collar.  Zeiss  makes  a  collar 
for  this  purpose. 

In  addition  to  the  Abbe  condenser,  there  should  be  another, 
placed  between  the  microscope  and  the  light.  For  this  purpose, 
the  large  condenser  from  a  projection  lantern  may  be  used.  For 


140  Methods  in  Plant  Histology 

magnifications  higher  than  100  diameters  still  another  condenser  will 
be  useful.  Place  it  between  the  last-named  condenser  and  the 
microscope. 

With  so  many  condensers,  the  heat  may  damage  preparations : 
so  place  between  the  last-named  condenser  and  the  one  next  the 
light  a  cooler  filled  with  water  or  a  solution  of  alum. 

With  all  these  accessories,  an  additional  iris  diaphragm  is  desir- 
able. Place  it  between  the  middle  one  of  the  three  condensers  and 
the  microscope,  but  quite  close  to  the  middle  condenser.  To  make 
an  efficient  adjustment  of  all  these  parts  requires  patience,  practice, 
and  judgment. 

It  will  save  time  and  patience  if  the  position  of  the  object  to  be 
photographed  be  marked  in  ink  on  the  slide  by  vertical  and  horizontal 
lines,  or  by  a  circle  drawn  around  it.  Even  with  these  lines,  it  is  none 
too  easy  to  get  the  object  into  the  desired  position  on  the  ground 
glass.  Remove  the  ground  glass  and  let  the  image  fall  on  a  piece 
of  white  cardboard  a  short  distance  back  of  the  camera.  If  the 
curtains  are  pulled  down,  the  position  of  the  object  in  the  field  and 
the  focusing  of  the  condensers  will  be  comparatively  easy. 

The  desirability  of  a  rigid,  straight,  and  accurate  optical  bed 
will  soon  be  realized.  If  one  is  intending  to  do  much  photomicro- 
graphic  work,  the  heavy,  graduated  optical  bed  is  almost  a  necessity. 
However,  if  time  is  no  object  and  patience  is  abundant,  good  photo- 
micrographs at  a  magnification  of  over  1,000  diameters  can  be  made 
with  no  apparatus  except  a  good  camera,  a  good  microscope,  and  a 
good  lamp. 

The  relative  positions  of  the  various  parts,  as  we  have  used  them 
in  making  the  illustrations  for  this  book,  are  indicated  below : 


-a     £ 

•2      2 

TZ  2-"  vi*i       *H  M     ^?    F^  fli    M^  ^_ 

o  -a    £    §    3s    ssa       66  ^ 

Some  data  which  may  be  helpful  will  be  found  in  the  legends  under 
some  of  the  photomicrographs. 


Photomicrographs  and  Lantern  Slides  141 

After  a  little  practice  the  student  will  read  with  profit  the  more 
extended  works  on  this  subject,  among  which  are  the  following: 
The  A,  B,  C  of  Photomicrography,  by  W.  H.  Walmsley  (Tennant  & 
Ward,  New  York);  Photomicrography,  ed.  by  J.  Spitta  (Scientific 
Press,  London,  England);  Lehrbuch  der  Microphotographie,  by 
Dr.  Richard  Neuhaus  (Harold  Bruhn,  Brunswick,  Germany). 

LANTERN  SLIDES 

Lantern  slides  are  now  so  universally  used  in  the  lecture-room 
that  every  teacher  should  be  able  to  make  them.  Three  general 
classes  of  lantern  slides,  as  far  as  the  technic  of  making  them  is 
concerned,  will  be  described:  (1)  lantern  slides  by  contact,  (2)  by 
reducing  or  enlarging,  and  (3)  by  copying  illustrations. 

1.  Lantern  Slides  by  Contact. — This  method  is  very  simple. 
Imagine  that  the  lantern-slide  plate  is  a  piece  of  printing-out  paper, 
and  proceed  just  as  in  making  a  print  on  paper.  Remember  that 
dust  on  the  negative  or  plate  causes  spots  in  the  print,  and  that  spots 
so  small  as  to  be  almost  unnoticeable  in  an  ordinary  print  will  be 
greatly  magnified  when  they  appear  on  the  screen.  Brush  both 
negative  and  plate  very  gently  with  a  soft  clean  brush  before  making 
the  print.  If  the  negative  is  3|X4£  inches,  it  can  be  placed  in  a 
printing  frame  of  that  size,  and  the  lantern  slide  placed  upon  it  with 
the  two  films  in  contact,  just  as  in  printing  paper.  If  there  is  no 
small  printing  frame,  use  a  4X5,  a  5X7,  or  even  an  8X10  frame. 
In  such  cases,  put  in  a  piece  of  clean  glass  free  from  scratches  or 
bubbles,  and  lay  the  negative  upon  it.  Lantern  slides  may  be 
printed  from  a  portion  of  a  4X5  or  some  larger  negative  by  simply 
placing  the  lantern-slide  plate  over  the  desired  spot.  Take  great 
care  not  to  scratch  the  negative. 

The  exposure  will  be  shorter  than  in  case  of  paper.  With  an 
average  negative  and  a  gas-mantle  lamp  at  a  distance  of  three  feet, 
try  an  exposure  of  2  seconds;  if  the  negative  is  weak,  shorten  the 
exposure;  if  strong,  lengthen  it. 

If  a  negative  is  uneven,  the  distance  from  the  light  may  be 
increased  so  as  to  lengthen  the  exposure  to  several  seconds,  thus  giving 
time  to  shade  the  weak  parts,  just  as  in  case  of  prints  on  paper. 


142  Methods  in  Plant  Histology 

If  a  negative  is  harsh  and  shows  too  much  contrast,  hold  it  closer  to 
the  light  and  shorten  the  exposure;  if  weak  and  lacking  in  contrast, 
hold  it  farther  away  and  increase  the  time  of  exposure. 

2.  Reducing  and  Enlarging. — If  a  slide  is  to  be  made  from  a  4  X  5 
or  larger  negative,  there  must  be  a  reduction.     A  camera  is  necessary. 
A  3|X4£  camera  is  large  enough.     If  any  larger  size  is  used,  the 
plate-holder  must  be  "kitted"  down  to  3|X4,  the  standard  size 
of  lantern  slides  in  America.     In  using  the  larger  cameras,  mark 
upon  the  ground  glass  the  exact  size  and  location  of  the  lantern- 
slide  plate.    Fasten  the  negative  in  some  convenient  place  where 
the  light  may  shine  through  it:    diffuse  daylight  is  good.     Then 
arrange  the  camera  just  as  in  taking  any  ordinary  picture.     The 
board  shown  in  Fig.  25  will  be  just  as  useful  in  making  lantern  slides 
as  in  making  photomicrographs.     At  one  end  of  the  board  fasten  a 
frame  which  will  hold  an  8  X 10  negative  and  also  hold  kits  for  smaller 
negatives  (Fig.  25,  B  and  C).     The  long  slot  in  the  board  will  allow 
the  camera  to  be  fastened  at  the  proper  distance.     If  buildings,trees, 
or  shadows  are  in  the  way,  tilt  the  board  so  as  to  have  a  clear  sky  for 
a  background. 

Be  very  careful  in  focusing;  it  is  best  to  examine,  with  a  pocket 
lens,  the  image  on  the  ground  glass.  In  general,  use  a  rather  small 
stop,  F  16  or  even  F  32.  If  reducing  from  an  average  5X7  negative, 
in  good  daylight,  with  an  F  16  stop,  try  3  or  4  seconds.  If  enlarging 
from  a  negative  somewhat  smaller  than  a  lantern  slide,  try  8  or  10 
seconds.  Other  things  being  equal,  the  best  lantern  slides  are  made 
by  reduction  from  larger  negatives  and  the  poorest  by  enlargement 
from  smaller  negatives. 

3.  Copying  Illustrations. — It  is  often  desirable  to  get  lantern 
slides  from  photographs,  maps,  or  pictures  in  books.     Here,  it  is 
necessary  to  make  a  negative  and  then  make  the  lantern  slide  from  the 
negative.     In  such  cases  make  a  3£  X4  negative  and  print  the  lantern 
slide  by  contact.     A  lantern-slide  plate  is  good  for  such  copying. 
The  exposure  will  depend  upon  the  light,  the  character  of  the  print, 
and  the  amount  of  reduction  or  enlargement.    Other  things  being 
equal,  the  exposure  will  always  be  longer  in  case  of  enlargement  than 
in  case  of  reduction.     If  an  average  5X7  photograph  is  to  be  copied 


Photomicrographs  and  Lantern  Slides  143 

in  good  diffuse  daylight,  with  an  F  16  stop  and  a  lantern-slide  plate, 
try  15  seconds. 

In  copying  maps  and  line  drawings,  where  dead  blacks  and 
pure  whites  are  desired,  expose  fully  and  overdevelop,  even  until 
the  image  shows  plainly  on  the  back  of  the  plate. 

It  is  not  necessary  to  furnish  formulae  for  developers  and  fixing 
solutions,  since  these  are  furnished  with  every  box  of  plates.  We 
have  found  the  Cramer  plates  very  satisfactory  for  all  kinds  of 
photographic  work.  The  firm  will  send  gratis  to  anyone  who  requests 
it  Cramer's  manual  on  Negative-Making  and  Formulas  (G.  Cramer 
Dry  Plate  Co.,  St.  Louis,  Missouri). 

To  the  formulae  in  common  use  may  be  added  one  by  Dr.  Land. 
It  is  good  for  general  work  and  gives  particularly  brilliant  results 
with  lantern  slides.  It  will  develop  an  underexposed  plate  when 
the  usual  developers  fail.  With  this  developer,  the  image  flashes 
into  sight  with  surprising  suddenness,  but  do  not  become  startled  and 
remove  the  slide  too  soon,  lest  you  fail  to  secure  details. 

Land's  Developer. — 

Hydrokinon 8  g. 

Metol 3g. 

Sodium  sulphite  (dry) 30  g. 

(60  g.  if  crystals  are  used) 

Sodium  carbonate  (dry) ,  30  g. 

(90  g.  if  crystals  are  used) 

Potassium  bromide 2  g. 

Water 1,000  c.c. 

Warm  Tones. — A  pyro-ammonia  developer  for  warm  tones  is 
recommended  in  Harrington's  Photographic  Journal  for  June,  1914: 

Metric         Apothecaries 

A.  Pyro 31  g.  (1  oz.) 

Sodium  sulphite  crystals 62  g.  (2  oz.) 

Citric  acid 2.6  g.  (40  g.) 

Water 237  c.c.     (    J  pint) 

B.  Ammonia 31  g.        (  1  oz.) 

Water 237  c.c.      ;    4  pint) 

C.  Ammonia  bromide 31  g.        (1  oz.) 

Water 237  c.c.      (    i  pint) 


144  Methods  in  Plant  Histology 

The  solutions  A,  B,  and  C  keep  well  separately,  but  not  when 
mixed.  When  wanted  for  immediate  use,  mix  A,  1 . 8  c.c.  (30  minims) , 
B,  3.7  c.c.  (60  minims),  and  C,  1 .8  c.c.  (30  minims),  and  add  30  c.c. 
(1  oz.)  of  water. 

A  correctly  exposed  plate  will  develop  in  about  2|  minutes,  and 
the  tone  should  be  a  warm  black.  Brown  tones  are  secured  by 
increasing  the  quantity  of  C,  while  A  and  B  remain  the  same. 

Reducing  Overexposed  Negatives  and  Lantern  Slides. — In  case 
of  overexposure,  the  negatives  or  lantern  slides  can  be  saved  by 
reducing.  The  reducing  solution  should  be  applied  as  soon  as  the 
negative  is  well  fixed  in  hypo.  If  a  negative  which  has  been  washed 
and  dried  is  to  be  reduced,  it  should  be  soaked  in  water  for  half  an 
hour  before  using  the  reducing  solution. 

The  following  is  a  good  solution  for  most  purposes : 

Metric     Apothecaries 

[Water 473  c.c.    (16  oz.) 

\Hyposulphite  of  soda 31  g.       (1  oz.) 

B  [Water 473  c.c.    (16  oz.) 

\Red  prussiate  of  potassium 31  g.       (  1  oz.) 

Solution  B  must  be  protected  from  the  light.  Cover  the  bottle 
with  black  paper  and  keep  it  in  the  dark  when  not  in  use. 

Mix  only  for  immediate  use  8  parts  of  A  to  1  of  B  and  use  in  rather 
subdued  light.  A  dark  room  is  not  necessary,  but  avoid  bright  light. 

When  the  negative  or  lantern  slide  becomes  satisfactory,  wash 
it  in  water  as  thoroughly  as  if  it  had  just  come  from  the  ordinary 
hypo  fixing  solution. 

Intensifying  Underexposed  Negatives  and  Lantern  Slides. — 
Even  if  a  negative  or  lantern  slide  has  been  considerably  overexposed, 
it  can  be  reduced  quite  satisfactorily;  if  much  underexposed,  little 
can  be  done  for  it;  if  only  slightly  underexposed,  it  may  be  greatly 
improved  by  the  following  solution: 

Metric  Apothecaries 

[Bichloride  of  mercury 2  g.  (31  gr.) 

A  |  Water 100  c.c.  (    4  oz.) 

[Bromide  of  potassium 2  g.  (31  gr.) 

[Sulphite  of  soda  crystals 10  g.  (154  gr.) 

\Water 100  c.c.  (    4  oz.) 


Photomicrographs  and  Lantern  Slides  145 

The  solutions  keep  indefinitely  and  may  be  used  three  or  four 
times. 

Apply  the  intensifier  after  fixing  in  hypo  and  washing  in  water. 
If  the  negative  or  slide  has  been  allowed  to  dry,  soak  it  in  water  for 
half  an  hour  before  intensifying. 

Place  the  negative  or  slide  in  A,  rocking  the  tray  as  in  developing, 
until  it  becomes  gray  or  even  white.  Wash  in  water  for  1  minute 
and  then  transfer  to  B  and  leave  until  the  dark  color  can  be  seen  on 
the  back  of  the  negative  or  slide.  Wash  in  water  as  thoroughly  as 
after  fixing  in  hypo. 

Some  use  a  saturated  aqueous  solution  of  the  bichloride  of  mer- 
cury, without  the  bromide  pf  potassium;  and,  instead  of  solution  B, 
use  water  to  which  ammonia  has  been  added — about  1  part  ammonia 
to  40  parts  water.  Excellent  sepia  tones  may  be  secured  in  this 
way.  Wash  well  in  water. 

After  the  plate  has  been  thoroughly  washed  in  water,  wipe  it 
gently  with  a  tuft  of  cotton.  The  cotton  must,  of  course,  be  thor- 
oughly wet;  it  is  better  to  hold  the  plate  under  a  stream  of  water 
while  wiping.  This  should  always  be  done  before  placing  a  negative  or 
slide  in  the  rack  to  dry,  after  a  washing  in  water. 

Toning  Lantern  Slides. — A  lantern  slide  may  sometimes  be  made 
more  effective  by  judicious  toning.  The  hints  given  here  merely 
introduce  the  student  to  the  possibilities  of  the  subject. 

Light  Sepia  to  Red  Tones. — Overexpose  up  to  four  or  five  times 
the  length  of  exposure  for  a  normal  slide  in  black  and  white;  develop 
thoroughly;  fix  and  wash  as  usual;  then  tone  in  the  following 
solution : 

Metric  Apothecaries 

A.  Potassium  ferricyanide 6  g.  (     90  gr.) 

Water 295  c.c.  (      10  oz.) 

B.  Copper  sulphate 7  g.  (    110  gr.) 

Potassium  citrate 65  g.  (1,000  gr.) 

Water 295  c.c.  (     10  oz.) 

(The  metric  and  U.S.  measures  are  practically  rather  than  arithmetically  equivalent. 

When  needed  for  use,  pour  some  of  A  into  an  equal  quantity  of 
B,  stirring  or  shaking  constantly.  Put  the  slide  into  the  solution  in  a 
tray  and  rock  just  as  if  developing  a  plate.  The  solution  is  a  strong 


146 


Methods  in  Plant  Histology 


reducer.  The  tone  should  change  from  black  to  warm,  then  to  sepia, 
and  may  finally  become  quite  red.  The  time  may  vary  from  1  to  20 
minutes,  according  to  the  density  of  the  slide  and  the  tone  desired. 
The  finished  product  must  not  be  too  dense,  for  a  slide,  toned  in  this 
way,  may  seem  rather  weak  and  yet  appear  surprisingly  strong 
on  the  screen. 

After  toning,  wash  in  water  for  about  20  minutes. 

Moonlight  Tints. — Some  excellent  formulae,  recommended  by 
Anderton,  will  be  of  service: 

Metric  Apothecaries 

Ferric  ammonia  citrate  (10  per  cent  solu- 
tion)    15  c.c.  (  £  oz.) 

Potassium  ferricyanide  (10  per  cent  solu- 
tion)    15  c.c.  (  5  oz.) 

Glacial  acetic  acid  (10  per  cent  solution) . .   148  c.c.  (5    oz.) 

The  following  gives  a  more  greenish-blue  tint: 

Metric        Apothecaries 

Uranium  nitrate  (10  per  cent  solu- 
tion in  water) 3.6  c.c.  (1  dram) 

Ferric  ammonia  citrate  (10  per  cent 
solution  in  water) 3 . 6  c.c  (1  dram) 

Potassium  ferricyanide  (10  per  cent 

solution  in  water) 7.2  c.c.  (2  drams) 

Nitric  acid  (10  per  cent  solution  in 
water) 7.2  c.c.  (2  drams) 

Both  solutions  intensify  considerably,  so  that  slides  to  be  toned 
should  be  rather  weak.  After  toning,  wash  20  minutes  in  water. 

Green  Tones. — 

A.  Potassium    bichromate    (10    per 

cent  solution  in  water) 60      drops 

Potassium   ferricyanide    (10    per 

cent  solution  in  water) 30      c.c.  (  1    oz.) 

Water 120      c.c.  (4    oz.) 

B.  Cobalt  chloride 3.9  g.  (60    gr.) 

Ferric  sulphate 3 . 9  g.  (60    gr.) 

Hydrochloric  acid 15      c.c.  (    3  oz.) 

Water 120      c.c.  (  4    oz.) 

Bleach  in  A,  wash  10  minutes  in  water,  tone  in  B,  and  then 
wash  20  minutes  in  water. 


Photomicrographs  and  Lantern  Slides  147 

Cleaning  Lantern  Slides. — Sometimes  a  slide  will  seem  perfectly 
clear,  just  as  it  comes  from  the  fixing  bath,  especially  from  an  acid 
fixing  bath;  usually,  however,  it  will  be  better  to  transfer  the  slide 
from  the  fixing  bath  to  a  weak  solution  of  acetic  acid — just  enough 
acid  to  give  the  solution  the  taste  of  weak  vinegar — and  then  rock 
for  a  minute  before  washing. 

The  following  clearing  fluid  may  be  used  in  the  same  way: 

Metric  Apothecaries 

Alum 20  g.  (1.3  gr.) 

Iron  sulphate 20  g.  (1.3  gr.) 

Citric  acid 20  g.  (1.3  gr.) 

Water 500  c.c.  (17      oz.) 

Coating  Lantern  Slides. — After  the  slide  has  become  thoroughly 
dry,  a  coat  of  balsam  or  shellac  will  add  much  to  its  brilliancy. 
Dilute  the  Canada  balsam  with  xylol  until  it  becomes  almost  as 
thin  as  water;  balance  the  slide  on  the  thumb  and  first,  second,  and 
third  fingers,  holding  it  as  level  as  possible;  pour  the  balsam  over 
it,  letting  the  balsam  flow  evenly  over  the  whole  surface;  then  tilt 
the  slide  and  pour  the  balsam  back  into  the  bottle.  Put  the  slide  in 
the  rack  to  dry. 

Mounting. — Add  a  suitable  mat  and  a  clean  lantern-slide  cover. 
Bind  the  two  together  with  a  lantern-slide  binding  strip.  Paste  on 
the  label,  or,  if  you  prefer,  put  the  label  on  the  mat  before  binding, 
so  as  to  have  it  protected  by  the  cover.  Lay  the  slide  down  so  that 
the  positions  are  just  as  they  were  in  the  original,  and  then  paste  the 
" thumb  mark"  in  the  lower  left-hand  corner. 


PART  II 


SPECIFIC  DIRECTIONS 

In  the  preceding  chapters  the  principles  and  methods  of  technic 
have  been  described  in  a  general  way.  It  is  difficult,  especially  for 
a  beginner,  to  apply  general  principles  to  specific  cases,  and,  besides, 
the  types  which  he  might  select  for  the  preparations  might  not  form 
a  symmetrical  collection.  Consequently,  a  series  of  forms  has  been 
selected  which  will  not  merely  serve  for  practice  in  microscopical 
technic,  but  will  also  furnish  the  student  with  preparations  for  a 
fairly  satisfactory  study  of  plant  structures  from  the  algae  up  to  the 
angiosperms.  It  is  not  at  all  our  purpose  to  discuss  general  morphol- 
ogy, but  rather  to  answer,  by  means  of  sketches  and  specific  direc- 
tions, the  multitudinous  questions  which  confront  the  instructor 
in  the  laboratory.  For  those  who  have  had  a  thorough  training 
in  general  morphology  the  following  suggestions  will  be  in  some 
degree  superfluous.  Those  who  are  beginning  the  study  of  minute 
plant  structure  are  referred  to  the  standard  textbooks  for  descrip- 
tions of  the  plants  mentioned  here. 

The  directions  for  collecting  and  growing  laboratory  material 
constitute  an  important  feature  of  this  part  of  the  book. 

With  a  few  exceptions,  the  order  in  which  the  forms  are  pre- 
sented is  that  given  in  Engler's  Syllabus  der .Pflanzenfamilien. 


151 


CHAPTER  XIII 


MYXOMYCETES  AND  SCHIZOPHYTES 
MYXOMYCETES 

With  the  exception  of  a  few  forms  like  Fuligo  (often  found  on 
oak  stumps  and  on  oak  bark  in  tanyards),  the  myxomycetes  are 
small,  and  are  usually  overlooked  by  collectors  (Fig.  26).  A  careful 
examination  of  rotting  logs  in  moist  woods  will  usually  reveal  an 
abundance  of  these  delicate  and  beautiful  organisms.  Various 

species  may  be  found  in  spring, 
summer,  and  autumn.  The  plas- 
modia  are  most  abundant  just 
after  a  warm  shower.  A  couple  of 
days  of  dry  weather  will  then 
bring  sporangia  in  abundance. 
The  specimens  should  be  pinned  to 
the  bottom  of  the  box  for  safe  car- 
rying. An  excellent  collecting-box 
can  be  made  from  an  ordinary 
paper  shoe-box.  On  the  bottom 
of  the  box  place  a  thin  piece  of  soft  pine,  or  a  piece  of  the  cor- 
rugated paper  so  commonly  used  in  packing;  or,  better  still,  a  sheet 
of  cork.  At  each  end  nail  in  a  piece  of  pine  half  an  inch  thick  and 
an  inch  high.  Upon  these  end  pieces  place  a  thin  piece  of  pine,  thus 
making  a  second  bottom,  which,  of  course,  should  not  be  fastened. 
A  second  pair  of  ends  with  a  third  pine  bottom  nailed  to  them  may 
rest  upon  the  second  bottom.  The  three  bottoms  will  give  a  con- 
siderable surface  upon  which  the  material  may  be  pinned.  For 
most  purposes,  the  specimens  are  simply  allowed  to  dry,  and  are 
then  fastened  with  glue  or  paste  to  the  bottom  of  a  small  box. 
Plasmodia  and  young  sporangia  may  be  fixed  in  chromo-acetic 
acid  or  Flemming's  fluid.  Sections  are  easily  cut  in  paraffin,  and 
should  not  be  more  than  5  n  in  thickness;  for  nuclear  details,  sections 

152 


FIG.  26. — Myxomycetes  growing  on 
rotten  wood:  A,  Hemiarcyria  rubiformis, 
X20;  B,  Stemonitis  ferruginea,  natural 
size;  C,  Trichia  varia,  Xli- 


Myxomycetes  and  Schizophytes  153 

should  not  be  thicker  than  2  ju  or  3  jw.  The  safranin,  gentian-violet, 
orange  combination  is  good  for  a  study  of  the  general  development 
and  for  some  cytological  features,  but  iron-alum  haematoxylin  is 
better  for  nuclear  details. 

Spores  of  most  myxomycetes  will  germinate  as  soon  as  they  are 
thoroughly  ripe,and,  during  the  first  year,  germination  is  more  prompt 
than  in  case  of  older  spores.  Fresh  spores  may  germinate  in  half 
an  hour;  the  time  may  extend  to  several  hours;  spores  two  or  three 
years  old  may  germinate  in  three  or  four  days,  or  may  not  germinate 
at  all.  We  have  never  succeeded  in  germinating  spores  which  were 
more  than  three  years  old.  The  longevity  is  doubtless  different 
in  different  species.  In  most  cases,  spores  will  germinate  in  water, 
if  they  will  germinate  at  all.  For  small  cultures,  the  hanging-drop 
method,  described  on  p.  73,  may  be  used. 

Plasmodia  may  be  raised  by  sowing  spores  on  moist,  rotten  bark 
or  wood  and  placing  the  culture  under  a  bell  jar,  where  the  moist, 
sultry  condition  favorable  to  their  growth  is  easily  imitated.  Plas- 
modia may  be  got  upon  the  slide  by  inclining  the  slide  at  an  angle  of 
about  15°,  with  one  end  of  the  slide  at  the  edge  of  the  plasmodium, 
and  allowing  water  to  flow  very  gently  down  from  the  upper  end  of 
the  slide  to  the  lower.  The  proper  flow  of  water  could  be  secured 
by  dropping  water  from  a  pipette,  but  a  less  tedious  plan  is  to  arrange 
a  siphon  so  as  to  secure  a  similar  current.  The  plasmodium  will 
creep  up  the  slide  against  the  current,  .furnishing  an  excellent  illus- 
tration of  rheotropism.  Enough  plasmodium  for  an  illustration  may 
be  formed  in  two  or  three  hours.  Examined  under  the  microscope, 
the  preparation  should  give  an  excellent  view  of  the  streaming 
movements  of  protoplasm. 

The  following  is  another  method  for  getting  the  plasmodia  upon 
the  slide :  Place  the  slides  upon  a  pane  of  glass  and  upon  each  slide 
place  a  small  piece  of  plasmodium-bearing  wood.  Cover  with  a  bell 
jar.  Wet  blotting  paper  or  a  small  dish  of  water  included  under  the 
jar  will  help  to  create  the  warm,  sultry  atmosphere  necessary.  The 
slides  may  be  covered  with  'the  plasmodium  in  a  few  hours.  Per- 
manent preparations  may  be  made  by  immersing  the  slide  in  chromo- 
acetic,  acid,  then  washing  and  staining  without  removing  the 


154 


Methods  in  Plant  Histology 


plasmodium  from  the  slide.  Acid  fuchsin  is  a  good  stain  for  bring- 
ing out  the  delicate  strands  of  the  plasmodium.  Iron-alum  haema- 
toxylin,  followed  by  acid  fuchsin  or  erythrosin,  brings  out  both  nuclei 
and  cytoplasmic  strands. 

Some  of  the  foregoing  methods  are  taken  from  an  article  by 
Professor  Howard  Ayers  in  the  January  and  February  (1898)  num- 
bers of  the  Journal  of  Applied  Microscopy.  Other  methods,  with 
directions  for  various  experiments,  are  given  in  the  same  article. 

SCHIZOPHYTES  (Fission  Plants) 

BACTERIA  (Schizomycetes,  Fission  Fungi) 

The  methods  of  modern  bacteriological  technic  are  so  numerous 

and   so   specialized   that   we   must    refer   to    laboratory   manuals 

for   instruction   in   this   subject.     The 

:     ,  >^t     \    ,it  ;  method  given  here  will  merely  enable 

-"">;     '    ;   ^.Jtp  the  student  to  study  the  form  and  size 

.r'X.      .-  :^''"v'  ^'^  of  those  bacteria  which  are  more  easily 

•";'_£     ;A       ^i7"-.^-  demonstrated. 

^^^\  ^  ^^i'..  Foul  water  at  the  outlets  of  sewers 

and  such  places  will  usually  afford  an 
•«  abundance  of  Coccus,  Bacillus,  Spiril- 

lum, and  Beggiatoa  forms.  Place  a  drop 
of  water  on  a  slide,  heat  it  gently  until 
the  water  evaporates,  then  stain  with 
fuchsin  or  methyl  violet,  dehydrate, 
clear  in  xylol,  and  mount  in  balsam 
(Fig.  27). 

The  hay  infusion  is  a  time-honored 
method  for  securing  bacteria  for  study. 
Pour  hot  water  on  a  handful  of  hay, 
and  filter  the  fluid  through  blotting 
paper.  Place  the  fluid  in  a  glass  dish, 
and  cover  with  a  piece  of  glass  to  keep 

out  the  dust.  When  the  fluid  begins  to  appear  turbid,  bacteria  will 
be  abundant.  The  active  movements  are  easily  observed  in  a 
mount  from  the  turbid  water.  As  the  bacteria  pass  into  the  resting 


PIG.  27.— Bacteria:  A,  Bacillus 
anthracis,  from  a  paraffin  section 
cut  from  the  liver  of  a  mouse;  fixed 
in  chromo-acetic  acid,  stained  in 
methyl  violet  and  Bismarck 
brown,  and  mounted  in  balsam; 
B,  Spirillum  sp.,  from  a  prepara- 
tion stained  in  fuchsin ;  C.  Staphy- 
lococcus  pyrogenes  aureus,  from  a 
preparation  stained  in  gentian- 
violet.  X535. 


Myxomycetes  and  Schizophytes  155 

condition,  they  form  a  scum  on  the  surface  of  the  water.  Usually, 
the  first  to  appear  is  a  somewhat  rod-shaped  form,  the  Bacterium 
termo  of  the  older  texts.  Spirillum  and  Coccus  forms  often  ap- 
pear later. 

Fine  preparations  may  be  obtained  by  inoculating  a  mouse  with 
Anthrax,  or  some  other  form,  and  then  cutting  paraffin  sections  of 
favorable  organs.  For  making  mounts  of  a  dangerous  form  like 
Anthrax,  secure  properly  fixed  material  from  a  bacteriologist. 
Gentian-violet  with  a  faint  Bismarck  brown  or  with  light  green  for  a 
background  makes  a  good  combination.  The  following  schedule 
gives  good  results  with  Anthrax  and  many  other  bacteria: 

1.  Gentian-violet,  5  minutes. 

2.  Rinse  in  water  a  few  seconds. 

3.  Gram's  solution  (iodine  1  g.,  potassium  iodide  2  g.,  water  300  c.c.) 
until  the  color  is  almost  or  quite  black;   this  will  generally  require 
1  or  2  minutes. 

4.  95  per  cent  alcohol  until  the  color  has  nearly  disappeared. 

5.  Rinse  in  water  and  examine.     If  the  bacteria  are  well  stained,  a 
counter-stain  may  be  added. 

6.  Light  green  or  erythrosin,  5  seconds;   or  Bismarck  brown,  5  or  10 
seconds. 

7.  95  and  100  per  cent  alcohol,  dehydrating  as  rapidly  as  possible.    Not 
more  than  5  or  10  seconds  can  usually  be  allowed. 

8.  Absolute  alcohol  and  xylol,  equal  parts,  3  or  4  seconds. 

9.  Xylol,  1  to  5  minutes. 
10.  Balsam. 

The  following  rapid  method  gives  fairly  good  results: 

1.  Place  on  a  clean  cover  a  drop  of  water  containing  the  bacteria  and 
dry  completely  in  a  flame  or  on  a  hot  plate. 

2.  Stain  2  to  5  minutes  in  gentian-violet  or  methyl  violet. 

3.  Rinse  quickly  in  water. 

4.  Dip  into  95  per  cent  alcohol  to  reduce  the  stain. 

5.  Remove  most  of  the  alcohol  by  touching  a  corner  of  the  cover  with 
filter  paper  and  then  dry  completely  by  passing  through  a  flame. 

6.  Mount  in  balsam. 

Leptothrix  may  often  be  obtained  by  scraping  the  inside  of  the 
cheek.     Beggiatoa,   one   of  the  sulphur  bacteria,   with  oscillating 


156  Methods  in  Plant  Histology 

movements  like   Oscillatoria,   is   often   found   in   foul   water.     Its 
presence  may  be  indicated  by  whitish  patches  on  the  bottom. 

The  Bacteria  are  the  only  plants  in  which  a  nucleus  has  not  been 
conclusively  demonstrated,  and  some  claim  that  a  nucleus  is  present 
even  in  Bacteria.  In  determining  the  presence  or  absence  of  a 
nucleus  in  Bacteria,  the  crude  method,  just  given,  would  be  of  no 
value,  and  even  the  most  critical  methods  of  the  bacteriologist,  who 
mounts  the  organisms  whole,  would  be  entitled  to  only  scant  con- 
sideration. The  presence  or  absence  of  a  nucleus  will  have  to  be 
determined  by  a  study  of  thin,  well-stained  sections  of  perfectly 
fixed  material. 

CYANOPHYCEAE.    BLUE-GREEN  ALGAE  (Schizophyceae  Fission  Algae) 

The  blue-green  algae  include  unicellular,  colonial,  and  filamentous 
forms.  They  occur  everywhere  in  damp  or  wet  places.  On  the 
vertical  faces  of  rocks  where  there  is  a  constant  dripping  of  water, 
brilliant  blue-green  forms  are  abundant.  In  the  Yellowstone 
National  Park  the  brilliant  coloring  of  the  rocks  is  due  in  large 
measure  to  members  of  this  group.  Many  forms  occur  as  brownish 
or  greenish  gelatinous  layers  on  damp  ground  or  upon  rocks,  or  even 
upon  damp  wooden  structures  in  greenhouses.  Other  forms  float 
freely  in  water. 

Oscillatoria. — For  most  purposes  it  is  best  to  study  Oscillatoria 
in  the  living  condition.  It  is  readily  found  in  watering-troughs,  in 
stagnant  water,  on  damp  earth,  and  in  other  habitats.  The  com- 
monest forms  have  a  deep  blue-green  or  brownish  color.  It  is  very 
easy  to  keep  Oscillatoria  all  the  year  in  the  laboratory.  Simply 
put  a  little  of  a  desirable  form  into  a  gallon  glass  jar  half  filled  with 
water.  By  adding  water  occasionally  to  compensate  for  evapora- 
tion, the  culture  should  keep  indefinitely.  In  a  jar  with  a  tightly 
fitting  -cover  we  have  kept  such  a  culture  for  years  without  renewing 
the  water. 

For  the  purposes  of  identification  and  herbarium  specimens  the 
material  may  simply  be  placed  on  a  slip  of  mica  and  allowed  to  dry. 
When  wanted  for  use,  add  a  drop  of  water  and  a  cover,  and  the  mount 
is  ready  for  examination.  After  the  examination  has  been  made, 


Myxomycetes  and  Schizophytes 


157 


remove  the  cover,  allow  the  preparation  to  dry,  and  then  return  it 
to  the  herbarium. 

Good  mounts  may  be  made,  especially  from  the  larger 
species,  by  the  Venetian  turpentine  method.  Fix  in  chromo- 
acetic  acid  and  stain  in  iron-haematoxylin  or  in  the  Magdala- 
red  and  anilin-blue  combination.  Either  stain  will  show  the  nuclei 
fairly  well. 

For  the  best  views  of  nuclei,  thin  sections  are  absolutely  necessary. 
Fix  in  Flemming's  weaker  solution,  get  the  material  into  paraffin 
by  the  gradual  processes  described  in  chap,  ix;  cut  2  to  5  ju  in  thick- 
ness, according  to  the  size  of  the  species;  stain  in  iron-haematoxylin, 
and  mount  in  balsam.  In  such  mounts,  the  scattered  condition  of 
the  material  as  it  appears  in  thin  sections  is  very  annoying.  As  soon 


PIG.  28. — Oscillatoria:    photomicrograph  from  a  paraffin  section  3  n  in 
thickness  and  stained  in  iron-alum  haematoxylin.      X373. 

as  the  material  is  thoroughly  washed  in  water,  arrange  it  so  that  the 
filaments  will  all  have  the  same  general  direction.  This  will  enable 
you  to  get  longitudinal  or  transverse  sections.  As  you  begin  with 
the  alcohols,  use  a  Petri  dish  and  lay  a  slide  over  the  material,  and 
keep  it  there  until  you  imbed  in  paraffin.  This  will  keep  the  fila- 
ments from  spreading  out  too  much,  and  you  will  be  able  to  get  as 
much  on  one  slide  as  you  would  be  likely  to  get  on  a  dozen  slides 
without  such  precaution. 

Oscillatoria,  as  it  appears  in  section,  is  shown  in  Fig.  28. 

Tolypothrix.— This  form  occurs  as  small  tufts,  either  floating 
in  stagnant  water  or  attached  to  plants  and  stones.  Some  species 
grow  upon  dam  ground.  It  furnishes  an  excellent  example  of  false 
branching  (Fig.  29).  Like  all  small  filamentous  algae,  it  may  be 
dried  on  mica  for  herbarium  purposes.  Venetian  turpentine  mounts 


158 


Methods  in  Plant  Histology 


k 


and  paraffin  sections  are  prepared  as  in  Oscillatoria.     Tolypothrix  is 
even  better  than  Oscillatoria  for  a  study  of  the  nucleus. 

Scytonema  is  a  similar  form  which  is  fairly  common.  It  is  often 
found  as  a  felt-like  covering  on  wet  rocks. 

In  staining  forms  like  Tolypothrix  and  Scytonema,  which  have  a 
thick  sheath,  take  care  not  to  obscure  the  cell  contents  by  staining 
the  sheath  too  deeply.  If  the  sheath  is  not 
stained  at  all,  you  may  not  be  able  to  see  the 
nature  of  the  false  branching.  Anilin  blue  is 
good  for  sections.  It  is  equally  good  for  Vene- 
tian turpentine  mounts,  but  is  likely  to  over- 
stain.  You  are  likely  to  get  good  mounts  with 
less  trouble  if  you  use  light  green  for  the  sheath. 
Nostoc. — Nostoc  is  a  cosmopolitan  form.  It 
occurs  on  damp  earth  or  floating  freely  in  water. 
Young  specimens  are  generally  in  the  form  of 
gelatinous  nodules,  but  in  older  specimens  the 
form  may  be  quite  various.  It  is  very  easy  to 
make  sections,  since  the  gelatinous  matrix  cuts 
well  and  holds  the  filaments  together.  Chromo- 
acetic  acid  is  a  good  fixing  agent.  Stains  which 
stain  the  gelatinous  matrix  make  the  prepara- 
tions look  untidy,  but  they  show  that  each 
filament  of  the  nodule  has  its  own  gelatinous 
sheath.  Small  nodules  may  be  stained  in  bulk 
and  be  got  into  Venetian  turpentine.  Crushed 
under  the  cover,  they  make  instructive  prepa- 
rations. 

Rivularia. — This  form  is  readily  found  on  the  underside  of  the 
leaves  of  water-lilies  (Nuphar,  Nymphaea,  etc.),  but  is  also  abundant 
on  submerged  leaves  and  stems  of  other  plants.  It  occurs  in  the 
form  of  translucent,  gelatinous  nodules  of  various  sizes.  Chromo- 
acetic  acid  gives  beautiful  preparations,  but  good  results  can  also 
be  secured  from  formalin  or  picric-acid  material. 

The  most  instructive  preparations  for  morphological  study  can 
be  obtained  by  the  Venetian  turpentine  method.  Stain  in  iron- 


FIG.  29.—  Tolypo- 
thrix, showing  "false 
branching":  h,  heter- 
ocyst ;  f ,  concave  cell ; 
6,  end  of  false  branch 
with  beginning  of  new 
sheath.  X620. 


Myxomycetes  and  Schizophytes  159 

haematoxylin  and  very  lightly  in  erythrosin,  the  latter  stain  being 
used  merely  to  outline  the  sheath.  When  ready  for  mounting, 
crush  a  small  nodule  under  a  cover-glass.  The  paraffin  method  is 
easily  applied,  since  the  gelatinous  matrix  keeps  the  filaments  in 
place.  Any  form  of  similar  habit  may  be  prepared  in  the  same  way. 

Gloeotrichia. — 
Gloeotrichia  (Fig.  30)', 
in  its  later  stages,  is  a 
free-floating  form.  In 
earlier  stages  it  is  at- 
tached to  various  sub- 
mersed aquatic  plants. 
Thenodules,  when 
young,  are  firm  like 
Nostoc,  but  as  they 
grow  older  and  larger 
they  become  hollow 
and  soft.  The  older 
forms  become  so  much 
dissociated  that  they 
lose  their  character- 
istic form  and  merely 
make  the  fixing  fluid 
look  turbid.  Allow  a 
drop  of  such  material 
to  spread  out  and  dry 
upon  a  slide  which  has 
been  slightly  smeared  with  albumen  fixative.  Leave  the  slide  in  95 
per  cent  alcohol  2  or  3  minutes  to  coagulate  the  albumen  fixative, 
and  then  stain  in  safranin.  If  the  background  appears  untidy,  stain 
for  24  hours,  or  longer;  you  can  then  extract  the  stain  from  the  back- 
ground, and  still  leave  the  long  spore  and  some  of  the  other  features 
of  the  filament  well  stained.  A  touch  of  cyanin  will  bring  out 
the  sheath.  Cyanin  and  erythrosin  is  a  good  combination  if  the 
material  is  clean.  The  firmer  nodules  may  be  treated  like  Nostoc  or 
Rivularia. 


FIG.  30. — Gloeotrichia:  photomicrograph  from  a  prepa- 
ration stained  in  cyanin  and  erythrosin;  negative  by 
Dr.  W.  J.  G.  Land. 


160 


Methods  in  Plant  Histology 


Wasserbltithe.— Many  genera  of  the  Cyanophyceae  occur  as 
scums,  often  iridescent,  on  the  surface  of  stagnant  or  quiet  water. 
Some  of  the  commonest  forms  are  Coelosphaerium  and  Anabaena 
(Fig.  31).  Some  of  the  Chlorophyceae  also  occur  as  Wasserbliithe. 
Where  the  material  is  very  abundant,  it  may  be  collected  by  simply 
skimming  it  off  with  a  wide-mouthed  bottle,  but  where  it  is  rather 

scarce  it  is  better  to  fil- 
ter the  water  through 
a  cloth  and  finally 
rinse  the  algae  off  into 
a  bottle.  Enough 
formalin  may  now  be 
added  to  the  water  in 
the  bottle  to  make  a 
3  per  cent  solution. 
The  material  may  be 
kept  here  indefinitely, 
but  after  a  few  hours 
it  is  ready  for  use.  If 
the  forms  are  small, 
like  Anabaena,  smear 
a  slide  lightly  with 
Mayer's  albumen  fixa- 
tive, as  if  for  paraffin 
sections,  add  a  drop 
of  the  material  and 
allow  it  to  dry  over 
night  or  for  24  hours; 
then  immerse  the  slide 
in  strong  alcohol  for  a  few  minutes,  and  then  proceed  with  the 
staining.  Cyanin  and  erythrosin  form  a  good  combination  for 
differentiating  the  granules.  Delafield's  haematoxylin,  used  alone, 
stains  some  granules  purple  and  others  red.  Iron-alum  haema- 
toxylin is  excellent  for  heterocysts.  If  the  forms  are  large  enough 
to  collapse  with  such  treatment  the  Venetian  turpentine  method  may 
be  employed. 


D 


E 


FIG.  31. —  Wasserbliithe:  A,  Coelosphaerium  Kiitzin- 
gianum;  B,  Anabaena  flos-aquae;  C,  Anabaena  gigantea; 
D  and  E,  a  heterocyst  and  a  spore  of  A.  gigantea  drawn 
from  paraffin  sections  stained  in  cyanin  and  erythrosin. 


Myxomycetes  and  Schizophytes  161 

If  it  is  desirable  to  make  paraffin  sections,  put  the  material,  drop 
by  drop,  on  a  piece  of  blotting  paper  until  an  appreciable  layer  of 
sediment  is  obtained.  Get  the  paper  with  its  material  into  paraffin 
by  the  usual  method,  taking  great  care  not  to  wash  the  algae  off. 
After  imbedding,  trim  away  the  paper  and  dip  the  block  in  melted 
paraffin.  Sections  can  now  be  cut  and  stained  in  the  usual  manner. 


CHAPTER  XIV 
CHLOROPHYCEAE.     GREEN  ALGAE 

Since  the  Chlorophyceae  furnish  our  best  illustrations  of  the 
evolution  of  the  plant  body,  the  origin  and  development  of  sex,  and 
also  the  beginning  of  alternation  of  generations,  they  occupy  a 
prominent  place  in  any  well-planned  course  in  the  morphology  of 
plants. 

They  are  found  in  both  fresh  and  salt  water,  but  are  most  abun- 
dant in  fresh  water.  The  ponds,  ditches,  and  rivers  of  any  locality 
will  yield  an  abundance  and  variety  both  of  the  unicellular  and  the 
multicellular  members  of  this  group.  Most  of  the  forms  are  inde- 
pendent, but  there  are  epiphytic,  endophytic,  and  saprophytic 
species.  The  larger  forms  and  those  which  grow  in  tufts  or  mats  are 
readily  recognized  in  the  field.  Many  of  the  smaller  forms  are 
attached  to  other  water  plants.  Drain  the  water  plants  and  then 
squeeze  them  over  a  bottle.  The  sediment  is  likely  to  contain  a 
variety  of  unicellular  and  other  small  algae. 

Many  of  the  genera  are  easily  kept  in  the  laboratory.  It  is  not 
necessary  to  have  very  large  aquaria.  Ordinary  glass  battery  jars 
holding  about  a  gallon  are  good  for  most  forms.  Jars  holding  two 
gallons  will  be  as  good  or  better.  For  some  cultures  which  are  to 
be  kept  for  a  long  time,  glass  jars  with  ground  tops  and  covers  are 
very  desirable.  Put  about  an  inch  of  pond  dirt  in  some,  clean  sand 
in  others,  and  in  still  others  use  a  gravel  bottom. 

When  possible,  use  the  water  in  which  the  algae  were  growing, 
since  very  few  take  kindly  -to  a  sudden  change  of  water.  If  the 
material  has  been  brought  to  the  laboratory  in  a  very  small  quantity 
of  water,  fill  the  jar  about  two-thirds  full  with  tap  water.  Let  the 
water  run  for  two  or  three  minutes  before  you  fill  the  jar,  since  the 
water  standing  in  the  pipes  is  injurious,  or  even  fatal,  to  most  algae. 
Add  water  occasionally,  only  a  little  at  a  time,  .to  compensate  for 
evaporation.  If  the  water  has  evaporated  until  the  jar  is  about 

162 


Chlorophyceae  163 

one-third  full  and  you  fill  it  nearly  to  the  top  with  tap  water,  you  are 
likely  to  kill  some  of  the  most  desirable  forms. 

It  is  a  mistake  to  put  too  much  material  into  a  jar.  A  wad  of 
Spirogyra  half  as  large  as  one's  finger  is  as  much  as  should  be  put 
into  a  gallon  jar.  As  it  grows  to  ten  or  twenty  times  that  amount 
it  is  not  necessary  to  keep  throwing  it  out,  since  it  will  gradually 
accommodate  itself  to  conditions.  However,  do  not  let  the  jar 
become  choked  with  the  material. 

Cultures  may  be  started  even  in  the  winter.  Bring  in  some  mud 
over  which  algae  were  growing  the  previous  summer  or  autumn; 
put  it  into  a  jar  and  fill  it  two-thirds  full  of  tap  water.  Also 
bring  in  sticks,  leaves,  and  stones  from  good  alga  localities  and 
put  them  into  jars  of  tap  water.  Cultures  may  be  started 
either  by  taking  mud  and  sticks  from  under  the  ice  or  by 
taking  them  from  places  which  have  entirely  dried  up  during  the 
summer  or  autumn.  A  few  such  jars  will  be  likely  to  yield  a 
variety  of  material. 

If  you  have  a  good  jar  of  Oedogonium,  or  some  other  desir- 
able form,  do  not  throw  it  out  if  the  alga  should  disappear. 
Remember  that  temporary  disappearances  occur  in  nature. 
Allow  the  culture  to  become  dry  and  then  set  it  aside  where 
it  will  be  protected  from  dust.  After  a  few  months,  pour  on 
tap  water  and  it  is  very  likely  that  you  will  soon  have  a  good 
jar  of  Oedogonium.  Many  algae  behave  similarly;  some,  like 
Volvox,  appear  for  a  short  time  and  then  disappear  for  a  long 
time;  some,  like  Cladophora,  may  last  the  whole  year  and  grow 
so  luxuriantly  that  the  excess  material  must  be  removed;  and 
some,  like  Ulothrix,  we  have  not  been  able  to  cultivate  at  all  in 
the  laboratory. 

Some  very  useful  hints  on  collecting  and  growing  fresh-water 
algae  for  class  work  will  be  found  in  an  article  by  Dr.  J.  A.  Nieuw- 
land  in  the  Midland  Naturalist,  1:85,  1909. 

Professor  Klebs  has  shown  that  various  phases  in  the  life  histories 
of  many  algae  and  fungi  may  be  produced  at  will.  By  utilizing  his 
results,  the  fruiting  condition  may  be  induced  in  many  of  the  common 
laboratory  types.  Knop's  solution  will  be  needed  in  most  cases. 


164  Methods  in  Plant  Histology 

A  stock  solution  which  can  be  diluted  as  required  may  be  made  as 
follows : 

Potassium  nitrate,  KN03 1  g. 

Magnesium  sulphate,  MgSC>4 1  g. 

Calcium  nitrate,  Ca(N03)2 3  g. 

Potassium  phosphate,  K2HP04 1  g. 

Dissolve  the  first,  second,  and  fourth  ingredients  in  1  liter  of 
distilled  water,  and  then  add  the  calcium  nitrate.  A  precipitate  of 
calcium  phosphate  will  be  formed.  For  practical  purposes  this  may 
be  called  a  0.6  per  cent  solution.  Whenever  a  dilute  solution  is 
made  from  the  stock  solution  the  bottle  must  be  shaken  thoroughly 
in  order  that  a  proper  amount  of  the  precipitate  may  be  included  in 
the  diluted  solution.  To  make  a  0 . 1  per  cent  solution,  add  5  liters  of 
distilled  water  to  1  liter  of  the  stock  solution ;  for  a  0 . 3  per  cent  solu- 
tion, add  1  liter  of  distilled  water  to  1  liter  of  the  stock  solution,  etc. 

The  addition  of  a  liter  of  a  0.2  per  cent  solution  to  4  or  5  liters 
of  water  will  often  produce  a  more  thrifty  growth.  Directions  for 
inducing  reproductive  phases  are  given  in  connection  with  the  various 
types.  With  a  good  supply  of  glass  jars,  plenty  of  Knop's  solution, 
a  reasonable  control  over  temperature,  and  the  teacher's  usual 
amount  of  patience,  most  laboratory  types  can  be  studied  in  the 
living  condition  at  all  seasons  of  the  year. 

Permanent  preparations  are  needed  to  show  details  which  are 
not  so  evident  in  the  fresh  material.  The  unicellular  and  filamentous 
members,  together  with  such  forms  as  Volvox,  are  best  prepared  by 
the  Venetian  turpentine  method.  The  structure  is  so  much  more 
complicated  than  in  the  Cyanophyceae  that  it  demands  far  more 
care  and  skill  to  make  good  preparations.  In  some  of  the  green 
algae,  like  Spirogyra  and  Closterium,  it  has  been  found  that  cell 
division  takes  place  most  abundantly  in  the  night;  mitotic  figures 
are  scarce  in  material  collected  in  the  daytime.  From  an  hour  before 
midnight  up  to  three  or  four  o'clock  in  the  morning  is  the  best  time, 
if  you  want  dividing  stages.  Chromo-acetic  acid  is  a  good  killing 
and  fixing  agent  for  the  whole  group.  Very  good  results  have  been 
obtained  by  adding  about  3  c.c.  of  1  per  cent  osmic  acid  to  100  c.c. 
of  chromo-acetic  acid  (Schaffner's  formula).  If  material  is  to  be 


Chlorophyceae  165 

sectioned,  Flemming's  weaker  solution,  or  this  solution  with  the  osmic 
acid  still  more  reduced,  is  likely  to  give  better  results  than  chromo- 
acetic  mixtures  without  any  osmic  acid.  A  formula  which  gives 
satisfactory  results  with  Spirogyra  may  cause  plasmolysis  with 
Cladophora.  A  few  filaments  should  be  placed  under  the  microscope 
in  the  fixing  agent,  and,  if  plasmolysis  occurs,  the  chromic  should  be 
weakened  or  the  acetic  strengthened  until  the  suitable  proportions 
are  determined.  This  is  a  slow  process,  but  difficult  forms  like 
Cladophora  and  Vaucheria  are  almost  sure  to  shrink  without  it. 
About  24  hours  in  any  of  the  chromic  series  and  a  24  hours'  washing 
in  water  will  be  sufficient  for  members  of  this  group.  Only  a  few 
of  the  most  commonly  studied  will  be  mentioned. 

With  Marine  Forms  use  sea-water  in  making  up  the  fixing  agents 
and  in  washing,  but  use  fresh  water  in  making  up  alcohols  and  for  the 
10  per  cent  glycerin. 

Volvox. — Volvox  is  found  in  ponds  and  ditches,  and  even  in 
shallow  puddles.  The  most  favorable  place  to  look  for  it  is  in  the 
deeper  ponds,  lagoons,  and  ditches  which  receive  an  abundance  of 
rain  water.  Volvox  is  often  associated  with  Lemna.  It  is  not  easy 
to  keep  an  abundance  of  Volvox  in  the  laboratory.  However,  when 
it  disappears,  do  not  throw  the  culture  out,  because  new  coenobia 
are  likely  to  develop  from  the  oospores. 

For  fixing,  use  chromo-acetic  acid  with  1  g.  chromic  acid  and 
2  c.c.  acetic  acid  to  200  c.c.  of  water.  The  addition  of  2  c.c.  of  Iper 
cent  osmic  acid  to  50  c.c.  of  the  solution  named  above  will  secure 
more  rapid  killing  and  fixing  and  will  bring  better  results  if  the 
material  is  to  be  sectioned. 

The  Venetian  turpentine  method  should  be  used  in  making 
mounts  of  the  whole  coenobium.  A  few  broken  bits  of  cover-glass 
should  be  placed  among  the  coenobia  to  prevent  any  pressure  by 
the  cover. 

For  paraffin  sections,  the  material,  preferably  in  sufficient 
abundance  to  make  a  layer  half  an  inch  deep  in  the  bottom  of  a  bottle 
as  large  as  one's  finger,  is  infiltrated  with  paraffin  in  the  usual  way. 
In  imbedding,  simply  pour  the  contents  of  the  bottle  out  so  as  to 
form  a  thin  layer  on  a  piece  of  glass.  If  a  dish  is  used,  the  paraffin 


166 


Methods  in  Plant  Histology 


cake  must  be  very  thin.     Fig.  32  shows  that  even  such  a  delicate 
organism  as  Volvox  can  be  imbedded  in  paraffin  without  shrinking. 
Dr.  Nieuwland  reports  that  Pandorina,  Eudorina,  and  Gonium, 
also  members  of  the  Volvocaceae,  are  commonly  found  in  summer  as 
constituents  of  the  green  scum  on  wallows  in  fields  where  pigs  are 
kept.     The  flagellate,  Euglena,  is  often  associated  with  these  genera. 
Pleurococcus. — This  form,  which  is  used  everywhere  as  a  labora- 
tory type  of  the  unicellular  green  algae,  is  found  on  the  bark  of  trees, 

where  it  is  more  abundant 
on  the  north  side  and 
near  the  ground.  It  is 
also  found  on  stones  and 
fences,  and  in  moist  situ- 
ations generally.  It  is 
easily  secured  in  nearly 
all  localities  and  at  all 


FIG.  32. —  Volvox:  photomicrograph  of  a  section 
stained  in  Delafleld's  haematoxylin ;  from  a  prepara- 
tion and  negative  by  Dr.  W.  J.  G.  Land. 


seasons. 

A  study  of  the  living 
material  is  sufficient  for 
any  general  course.  The 
bright-green  cells,scraped 
off  and  mounted  in  a 
drop  of  water,  show  the 
rather  thick  wall,  the 
chromatophores,  and 
usually  the  nucleus.  A 
drop  of  iodine  will  bring  out  the  nucleus,  if  it  does  not  show 
already,  and  will  also  stain  the  pyrenoid,  if  the  cell  contains  one. 
A  mount  in  Venetian  turpentine,  stained  in  Magdala  red  and  anilin 
blue,  shows  the  nucleus  very  clearly. 

Scenedesmus. — Scenedesmus  (Fig.  33)  is  found  everywhere  as  a 
regular  constituent  of  the  fresh-water  plankton.  It  is  more  abundant 
in  stagnant  water.  It  often  appears  in  considerable  quantity  in 
laboratory  cultures.  It  may  be  kept  for  years  in  a  tightly  closed 
glass  jar  without  renewing  the  water,  the  lid  being  removed  only 
when  material  is  needed. 


Chlorophyceae 


167 


The  form  is  so  small  that  in  living  material  little  more  than  the 
general  form  can  be  distinguished.  Excellent  mounts  are  easily  and 
quickly  made.  Smear  a  very  thin  layer  of  albumen  fixative  upon  the 
slide,  and  add  a  drop  of  water  containing  the  Scenedesmus.  The 
drop  may  be  inverted  for  1  or  2  minutes  over  the  fumes  of  1  per  cent 
osmic  acid.  No  washing  is  necessary,  and  good  mounts  may  be 
made  without  any  fixing  whatever.  Allow  the  drop  to  dry  com- 
pletely. It  is  better  to  leave  it  for  24  hours  before  proceeding.  The 


FIG.  33. — Scenedesmus:  photomicrograph  from  a  preparation  by  Dr.  Yama- 
nouchi,  mounted  whole  and  stained  as  described  in  the  text;  Cramer  contrast 
plate;  4  mm.  objective;  ocular  X4;  yellowish-green  filter;  camera  bellows,  1 
meter;  arc  light;  exposure,  6  seconds.  X675. 

usual  difficulty  with  this  form,  and  with  many  others,  is  that  the 
background  stains  and  so  makes  the  mounts  untidy.  The  following 
method  by  Yamanouchi  will  produce  beautiful  preparations  (Fig.  33) : 

1.  Dry  on  the  slide,  24  hours. 

2.  10  per  cent  alcohol  over  night  to  remove  chlorophyll. 

3.  Safranin  (alcoholic),  4  days. 

4.  Water,  5  minutes. 

5.  Aqueous  gentian-violet,  2  days. 

6.  Water,  a  few  seconds. 

7.  Orange  G,  aqueous,  3  minutes. 


168 


Methods  in  Plant  Histology 


8.  95  per  cent  alcohol,  a  few  seconds. 

9.  Absolute  alcohol,  1  minute. 

10.  Clove  oil,  until  the  stain  is  satisfactory.    Different  collections  of 
Scenedesmus  stain  very  differently,  but  the  time  in  clove  oil  is  likely 
to  be  long,  even  as  long  as  6  hours. 

11.  Xylol,  5  minutes. 

12.  Mount  in  balsam. 

Hydrodictyon. — This  is  popularly  known  as  the  "water-net." 
Hydrodidyon  is  found  floating  or  suspended  in  ponds,  lakes,  or  slow 
streams.  The  young  nets  are  formed  within  the  segments  of  the 
older  nets.  Examine  segments  4  or  5  mm.  in 
length  for  the  formation  of  young  nets.  The 
old  nets  may  reach  a  length  of  10  cm. 
Cultures  are  easily  kept  in  the  laboratory. 
If  material  which  has  been  growing  in  a  0.5 
to  1  per  cent  Knop's  solution  be  brought  into 
tap  water  or  pond  water,  zoospore  formation 
may  begin  within  24  hours.  Nets  brought 
from  the  nutrient  solution  into  a  1  to  4  per 
cent  cane-sugar  solution  produce  zoospores  for 
a  few  days. 

Nets  of  all  sizes  should  be  selected  for 
study.  The  segments  are  coenocytic,  and  the 
nuclei  of  the  older  segments  are  hard  to 
differentiate,  except  in  stained  preparations.  Only  one  nucleus  will 
be  found  in  the  young  segments,  but  in  the  older  segments  the  nuclei 
become  very  numerous. 

For  fixing,  use  the  chromo-acetic  solution  recommended  for 
Vaucheria.  The  Venetian  turpentine  method  should  be  used  for 
mounting  entire  young  nets  or  entire  segments  of  older  nets.  Mag- 
dala  red  with  a  rather  light  stain  in  anilin  blue  brings  out  the  nuclei 
and  pyrenoids.  For  young  nets  inside  the  old  segments,  the  blue 
should  be  a  little  deeper.  Use  fine  scissors  very  freely:  teasing 
with  needles  is  ruinous.  Hydrodictyon  is  easily  imbedded  and  cut. 
Iron-haematoxylin  or.  the  safranin,  gentian-violet  combination  are 
best  for  paraffin  sections  (Fig.  34). 


FIG.  34.— Hy d rod i c- 
tyon:  zoospores  becoming 
arranged  into  a  new  net 
inside  an  old  segment; 
Venetian  turpentine 
method.  X125. 


Chlorophyceae  169 

Ulothrix. — Where  the  problem  of  the  origin  and  evolution  of  sex 
is  studied,  Ulothrix  is  an  indispensable  type.  Ulothrix  zonata  is 
found  in  springs,  brooks,  and  rivers,  occurring  in  bright-green  masses 
attached  to  stones  in  riffles,  especially  in  sunny  places.  It  is  abun- 
dant on  stones  and  piles  along  the  beaches  of  lakes.  Another  species 
is  found  in  stagnant  ponds,  ditches,  and  even  in  watering-troughs 
and  rain-barrels.  It  is  difficult  to  keep  in  the  laboratory  the  forms 
which  are  found  in  rapidly  flowing  water.  However,  if  they  are 
brought  in  still  attached  to  stones  and  placed  under  a  stream  of  tap 
water,  they  may  live  for  a  couple  of  weeks  and  may  produce  zoospores 
every  morning.  The  production  of  zoospores  may  continue  for  a 
few  days,  if  the  material  is  merely  put  into  a  jar  of  water;  in  a  2  to  4 
per  cent  cane-sugar  solution  the  production  of  zoospores  continues 
a  little  longer. 

While  the  most  instructive  study  demands  living  material, 
some  details  are  more  easily  seen  in  stained  preparations.  Fix  in 
chromo-acetic  acid  and  use  the  Venetian  turpentine  method.  Stain 
in  iron -alum  haematoxylin.  It  is  a  good  plan  to  stain  some  of  the 
material  in  Magdala  red  and  anilin  blue.  When  mounting,  some 
material  from  each  lot  can  be  used  for  every  preparation.  The 
iron-haematoxylin  will  give  the  best  views  of  the  nucleus;  the  anilin 
blue  will  stain  the  chromatophore  and  cell  well.  In  general,  it  is  a 
good  plan  to  put  upon  the  same  slide  material  prepared  in  various 
ways.  A  single  preparation  will  then  afford  a  rather  complete  study. 

Oedogonium. — This  form  is  attached  when  young,  but  most 
species-  float  freely  when  they  are  older.  Most  species  are  found 
in  quiet  waters,  especially  in  ponds  and  ditches.  The  floating 
masses  bear  some  resemblance  to  Spirogyra,  but  are  not  so  slippery. 
The  best  fruiting  material  is  often  found  attached  to  twigs,  rushes, 
and  various  plants,  where,  to  the  naked  eye,  it  forms  only  a  fuzzy 
covering  rather  than  a  dense  mat. 

In  studying  Oedogonium  diplandrum,  Klebs  found  that  a  change 
from  a  lower  to  a  higher  temperature  would  induce  the  production 
of  zoospores.  A  culture  which  had  been  kept  in  a  cold  room  with 
a  temperature  varying  from  6°  to  0°  C.,  when  brought  into  a  warmer 
room  with  a  temperature  varying  from  12°  to  16°  C.,  produced  an 


170 


Methods  in  Plant  Histology 


abundance  of  zoospores  within  two  days.  Light  does  not  seem  to 
have  any  influence  upon  the  formation  of  zoospores  in  this  species, 
but  light  is  necessary  for  the  formation  of  antheridia  and  oogonia. 
Any  culture  solutions  must  be  very  weak.  Sterile  material  sometimes 
fruits  when  brought  into  the  laboratory  and  placed  in  open  jars  with 
plenty  of  water  and  not  too  much  light. 

Fix  in  chromo-acetic  acid  and  use  the  Venetian  turpentine 
method.  Iron-haematoxylin  is  good  for  antheridia  and  also  for 
nuclei  and  pyrenoids,  but  anilin  blue  is  better  for 
caps  and  cell  walls  and  for  some  of  the  cell  con- 
tents. It  is  better  to  stain  material  in  both  ways 
and  then  put  some  from  each  lot  on  every  slide 
(Fig.  35). 

Coleochaete. — Coleochaete  is  epiphytic  upon 
the  stems  and  leaves  of  submerged  plants.  C. 
scutata,  which  is  the  most  common  species,  has  a 
flat,  orbicular  thallus  generally  less  than  1  mm. 
in  diameter.  C.  pulvinata  has  a  hemispherical 
thallus  and  might  be  mistaken  for  Rivularia,  un- 
less examined  with  a  lens. 

For  most  purposes  it  is  better  to  mount  the 
whole  plant.  Complete  the  staining  before  trying 
to  remove  the  Coleochaete  from  its  host.  Dela- 
field's  haematoxylin  is  a  good  stain.  Test  the 
staining  by  removing  single  specimens  and  examin- 
ing them  under  the  microscope.  When  the  stain- 
ing is  satisfactory,  wash  thoroughly  in  water  and 
transfer  to  10  per  cent  glycerin  and  follow  the  Venetian  turpen- 
tine method.  When  the  turpentine  is  thick  enough  for  mounting, 
remove  the  plants  from  the  stem  or  leaf  and  make  the  preparations. 
The  plants  may  be  removed  before  fixing  or  at  any  stage  in  the 
process,  but  they  are  so  small  that  great  care  must  be  taken  not  to 
lose  them  when  changing  solutions. 

Sections  are  easily  cut  and,  especially  in  forms  with  a  flat  thallus, 
show  features  which  might  escape  if  one  depended  entirely  upon 
plants  mounted  whole.  Cut  out  small  pieces  of  leaf  or  stem 


FIG.  35. — Oedogo- 
ntum:  egg  shortly 
after  fertilization, 
showing  nucleus, 
pyrenoids,  and  food 
material.  X465. 


Chlorophyceae  171 

abundantly  covered  with  Coleochaete,  imbed  in  paraffin,  and  cut 
host  and  guest  together. 

Diatoms. — Living  diatoms  are  often  found  clinging  in  great 
numbers  to  filamentous  algae,  or  forming  gelatinous  masses  on  vari- 
ous submerged  plants.  Cladophora  is  frequently  covered  with 
Cocconeis,  an  elliptically  shaped  diatom ;  Vaucheria  is  often  covered 
with  small  forms.  Other  algae  will  pay  for  examination,  especially 
if  they  look  brown.  If  stones  in  the  water  have  a  brown,  slippery 
coating,  you  can  be  sure  of  diatoms.  Sometimes  the  brown  coat- 
ing on  sticks  and  stones  is  so  abundant  that  it  streams  out  with  the 
current.  If  rushes  and  stems  of  water  plants  have  a  brown,  gelati- 
nous coating,  you  are  likely,  to  find  millions  of  specimens  of  the  same 
diatom.  The  surface  mud  of  a  pond,  ditch,  or  lagoon  will  always 
yield  some  diatoms.  They  can  be  made  to  come  out  from  the  mud 
by  putting  a  black  paper  around  the  jar  and  letting  direct  sunlight 
fall  upon  the  surface  of  the  water.  The  diatoms,  in  a  day  or  even 
less,  will  come  to  the  top  in  a  scum  which  can  be  easily  secured. 

Fresh-water  diatoms  appear  in  greatest  abundance  in  spring, 
are  comparatively  scarce  in  summer,  and  reappear  in  autumn, 
though  not  so  abundantly  as  in  the  spring. 

Marine  forms  can  be  secured  by  scraping  barnacles,  oyster  shells, 
and  other  shells.  The  big  Strombus  shell  from  the  West  Indies,  which 
we  use  to  keep  the  door  open,  will  yield  a  good  collection  if  you  get 
it  before  it  is  cleaned. 

The  silicious  shells  of  diatoms  are  among  the  most  beautiful 
objects  which  could  be  examined  with  the  microscope  (Fig.  36). 
To  obtain  perfectly  clean  mounts  requires  considerable  time  and 
patience,  but  when  the  material  is  once  cleaned,  preparations  may 
be  made  at  any  time  with  very  little  trouble.  Diatom  enthusiasts 
have  devised  numerous  methods  for  cleaning  them,  and  separating 
the  various  forms  from  each  other,  but  we  shall  give  here  only  a  few 
simple,  practical  methods. 

Material  for  mounts  of  frustules  of  living  forms  "may  be  obtained 
by  skimming  off  the  brownish  scum  found  on  ponds,  by  squeezing 
out  water  weeds,  by  scraping  sticks  and  stones  which  are  covered  at 
high  water,  or  from  the  mud  of  filter  beds  and  pumping-worka,  or 


172 


Methods  in  Plant  Histology 


in  other  places.  The  material  is  put  in  a  dish  of  water,  and  after  it 
has  settled  the  water  is  decanted.  This  is  repeated  until  the  water 
will  clear  in  about  half  an  hour.  The  sediment  is  then  treated  with 
an  equal  bulk  of  sulphuric  acid,  after  which  bichromate  of  potash 
is  added  until  all  action  ceases.  After  a  couple  of  hours  the  acid  is 


FIG.  36. — Diatoms:  diatomaceous  earth  from  Cherryfleld,  Maine,  showing  the 
great  variety  of  forms  usually  found  in  such  material ;  photomicrograph  from  a  prepara- 
tion by  Rev.  E.  L.  Little.  X400. 

washed  out.  To  separate  the  diatoms,  place  the  sediment  in  a  glass 
dish  with  water,  and  when  the  water  becomes  clear  give  the  dish  a 
slight  rotary  motion.  This  will  bring  the  diatoms  to  the  top,  when 
they  may  be  removed  with  a  pipette  and  placed  in  alcohol.  To 
mount,  place  a  number  in  distilled  water,  evaporate  a  few  drops  of 


Chlorophyceae  173 

the  mixture  on  a  cover-glass,  which  is  then  mounted  on  a  slide  in 
balsam."1 

Many  scouring  soaps  and  silver  polishes  contain  large  quantities 
of  fossil  diatoms,  and  the  diatomaceous  earths  are  particularly  rich. 
Break  up  a  small  lump  of  such  material  and  boil  it  in  hydrochloric 
acid.  A  test-tube  is  very  convenient  for  this  process.  Let  the 
diatoms  settle,  pour  off  the  acid,  and  then  wash  in  water.  As  soon 
as  the  diatoms  settle,  the  water  should  be  poured  off.  The  washing 
should  be  continued  until  the  hydrochloric  acid  has  been  removed. 
When  the  washing  is  complete,  pour  on  a  little  absolute  alcohol,  and 
after  a  few  minutes  pour  off  the  alcohol  and  add  equal  parts  of 
turpentine  and  carbolic  acid.  The  material  will  keep  indefinitely  in 
this  condition,  and  may  be  mounted  in  balsam  at  any  time.  In 
making  a  mount,  put  a  little  of  the  material  on  a  slide  and  allow  it 
to  become  dry,  or  nearly  dry,  and  then  add  the  balsam  and  cover.  If 
the  balsam  should  be  added  too  soon,  the  diatoms  are  likely  to  move 
to  the  edge  of  the  cover. 

To  show  the  cell  contents,  diatoms  must  be  fixed  and  stained. 
If  they  are  clinging  to  filamentous  algae,  the  algae  with  the  diatoms 
attached  should  be  put  into  chromo-acetic  acid  (24  hours)  and  then 
washed  in  water  for  24  hours.  Stain  in  iron-haematoxylin  and 
proceed  by  the  Venetian  turpentine  method.  When  ready  for 
mounting,  the  diatoms  can  be  scraped  off  from  the  algae  or  other 
substratum.  Other  stains  may  be  used. 

When  the  material  is  in  gelatinous  masses  it  may  be  fixed  in 
chromo-acetic  acid,  with  or  without  a  little  osmic  acid,  and  imbedded 
in  paraffin.  There  will,  of  course,  be  some  difficulty  in  cutting, 
but  the  knife  often  breaks  the  frustules  very  cleanly,  so  that  good 
sections  may  be  secured.  It  might  be  worth  while  to  try  a  weak 
solution  of  hydrofluoric  acid  to  dissolve  the  silicious  shells. 

Desmids.— The  desmids  are  unicellular,  free-floating  or  sus- 
pended algae.  They  are  much  more  abundant  in  soft  water  than  in 
hard.  Deep  pools,  quiet  ponds,  and  quiet  margins  of  small  lakes 
are  good  collecting-grounds.  Collections  of  other  fresh-water  algae 

i  From  a  review  of  Dr.  Wood's  paper  on  "Diatoms,"  Journal  of  Applied  Microscopy, 
March,  1899. 


174  Methods  in  Plant  Histology 

often  contain  some  desmids.  It  frequently  happens  that  a  single 
desirable  desmid  appears  during  examination  of  field  collections. 
In  such  a  case,  remove  it  with  a  fine  pipette,  and  get  it  into  a  drop  of 
water  on  a  clean  slide,  invert  it  over  a  bottle  of  1  per  cent  osmic  acid 
for  2  minutes,  leave  the  slide  exposed  to  the  air  until  almost  all  the 
water  has  evaporated,  and  then  add  a  drop  of  10  per  cent  glycerin- 
In  a  few  hours  (6  to  24)  put  on  a  cover  and  seal.  It  requires  more 
time,  care,  and  patience  than  it  is  worth  to  attempt  staining  in  such 
a  case. 

Sometimes  desmids  occur  in  great  abundance.     They  may  then 
be  treated  like  the  filamentous  algae,  except  that  more  care  must 
be  taken  not  to  lose  them  when  changing  fluids.     The  Venetian 
turpentine  method,  with  Magdala  red 
and  anilin  blue,   will   give   beautiful 
preparations.     A  deep  stain  with  Mag- 
dala red  and  a  rather  light  stain  with 
anilin  blue  is  better  for  the  pyrenoids 
and  nucleus,  while  a  light  stain  in  the 
red  and  a  deep  stain  in  blue  is  better 
PIG.  37.— Zygnema.-  photomicro-     for  the   chromatophores.     When   the 

graph  from  a  preparation  stained  .    .    .  fF-     •         .  i  i  i 

in   iron-alum    haematoxyiin    and     material  is  sufficiently  abundant, 

moulted   in    Venetian   turpentine.       paraffin     sections     may     be     made     as 

directed  for  Volvox. 

Lutman  has  found  that  Closterium  divides  at  night.  If  mitotic 
figures  are  wanted  they  are  more  likely  to  be  obtained  if  the  material 
is  fixed  about  midnight. 

Zygnema. — Zygnema  is  one  of  the  commonest  algae  of  the  ponds, 
swamps,  and  ditches.  The  mats  are  very  slippery  to  the  touch. 
In  the  field  it  resembles  Spirogyra,  but  is  distinguished  by  the  two 
characteristic  chromatophores  which  are  readily  seen  with  a  good 
pocket  lens.  Sometimes  conjugation  can  be  induced  by  bringing 
the  material  into  the  laboratory  and  placing  it  in  open  jars  with 
plenty  of  water  and  not  too  much  light. 

Iron-haematoxylin  is  a  good  stain  for  conjugating  material.  The 
stain  should  be  extracted  until  the  four  chromatophores  become 
distinct.  The  nuclei  are  small  and  inconspicuous.  The  chromato- 


Chlorophyceae 


175 


phores  do  not  stain  as  readily  as  those  of  Spirogyra,  and  consequently 
it  is  necessary  to  use  stronger  stains  or  more  prolonged  periods.  Use 
the  Venetian  turpentine  method  (Fig.  37). 

For  a  detailed  study,  imbed  in  paraffin  and  cut  thin  sections. 
After  washing  in  water,  arrange  the  filaments  so  that  most  of  them 
will  have  the  same  general  direction;  then,  in  running  up  through 
the  alcohols,  keep  the  filaments  from  spreading  too  much  by  placing 
a  slide  on  the  material.  After  imbedding,  the  material  can  be  cut 
into  blocks  about  a  centimeter  square.  If  sections  thinner  than  5  ju 
are  wanted,  cut  out  smaller  paraffin  blocks. 

Spirogyra. — Probably  no  alga  has  been  more  studied  by  pupils, 
teachers,  and  investigators  than  Spirogyra  (Fig.  38).  Nearly  all  of 


pIG.  38 Spirogyra:    photomicrograph  of  zygospores  (elliptical  in  outline)  formed 

by  lateral  conjugation;  the  spore  with  circular  outline  has  been  formed  without  conjuga- 
tion and  is  called  a  "  parthenogenetic "  spore  or  an  "  azygospore " ;  fixed  in  5  per  cent 
formalin  and  stained  in  iron-alum  haematoxylin.  X275. 

the  numerous  species  belong  to  the  low,  quiet  waters  of  ponds  and 
ditches,  where  they  often  form  large,  flocculent  green  mats  nearly 
covering  the  surface  of  the  water.  A  few  species  occur  in  running 
water.  The  mats  are  very  slippery  to  the  touch— a  character  which 
assists  in  recognizing  the  genus  in  the  field.  In  the  larger  species 
the  characteristic  spiral  chromatophore  can  be  seen  with  a  good 
pocket  lens,  thus  completing  the  identification,  as  far  as  the  genus  is 
concerned.  Mats  in  which  zygospores  have  been  formed  are  likely 


176  Methods  in  Plant  Histology 

to  show  a  pale,  or  even  a  brownish,  color,  due  to  the  brownish  walls 
of  the  zygospores.  This  color,  however,  is  not  always,  or  even 
usually,  due  to  zygospores,  but  is  more  often  due  to  the  death  and 
degeneration  of  the  plants.  Mats  in  early  stages  of  conjugation  and 
those  with  young  zygospores  show  as  bright  a  green  as  vigorously 
growing  material. 

Spirogyra  is  not  easy  to  keep  in  the  laboratory.  The  small 
species  keep  better  than  the  larger  ones.  Put  only  a  small  amount  of 
the  material  in  a  jar  and  use  rain  water.  If  it  is  necessary  to  use  tap 
water,  let  the  water  run  for  a  minute  before  taking  the  water  for  the 
culture.  Most  metals  are  poisonous  to  Spirogyra,  even  the  small 
amount  taken  up  by  the  water  while  standing  in  the  water  pipe  being 
detrimental. 

The  species  found  in  running  water  will  usually  conjugate  within 
a  week  when  brought  into  the  laboratory  and  placed  in  rainwater  or 
tap  water.  Species  belonging  to  quiet  waters,  when  brought  into 
the  laboratory  and  placed  in  a  0.2  per  cent  Knop's  solution,  are 
likely  to  undergo  rapid  cell  division  and  growth.  After  the  alga 
has  remained  in  such  a  culture  for  a  few  days  or  for  a  week,  conjuga- 
tion may  be  induced  by  transferring  to  rain  water  or  tap  water,  and 
keeping  the  culture  in  bright  sunlight.  Conjugation  may  begin 
within  3  or  4  days.  Variations  in  temperature  between  1°  and  15°  C. 
have  little  influence  upon  conjugation. 

The  following  is  a  good  fixing  agent  for  most  species  of  Spirogyra: 

Chromic  acid 1  g. 

Glacial  acetic  acid 4  c.c. 

Water 400  c.c. 

Fix  24  hours  and  wash  24  hours  in  running  water.  Use  the  Vene- 
tian turpentine  method.  With  Magdala.  red  and  anilin  blue  the 
most  beautiful  preparations  are  rather  easily  obtained,  the  spiral 
chromatophore  taking  the  blue  and  its  pyrenoids  the  red.  If  the 
material  contains  figures,  stain  in  iron-haematoxylin.  This  will  stain 
the  figures,  but  will  hardly  touch  the  chromatophore  or  cell  wall, 
thus  allowing  an  unobstructed  view  of  the  figures.  While  figures 
occur  occasionally  in  the  daytime,  collect  your  material  at  night, 
preferably  near  midnight. 

Spirogyra  is  easily  imbedded  and  cut. 


Chlorophyceae  177 

Vaucheria.— This  form  can  always  be  obtained  in  greenhouses, 
especially  in  the  fernery,  where  it  forms  a  green  felt  on  the  pots. 
The  greenhouse  form  is  likely  to  be  Vaucheria  sessilis.  Another 
species,  V.  geminata,  is  very  common  in  the  spring,  when  it  may 
be  found  in  ponds  and  ditches  (Fig.  39).  Vaucheria  is  also  found  in 
running  water,  but  in  this  situation  is  almost  certain  to  be  sterile. 
In  the  vicinity  of  Chicago,  V.  geminata  appears  late  in  March  or 
early  in  April  and  within  a  few  weeks  begins  to  fruit  abundantly. 
The  fruiting  continues  for  4  to  8  weeks,  and  then  the  alga  may 
disappear  until  later  in  the  season,  when  some  of  the  oospores 
germinate. 

Vaucheria  sessilis  is  found  at  all  seasons  in  the  greenhouses,  but 
it  is  usually  in  the  vegetative  condition.  Klebs  found  that  the 
formation  of  oogonia  and 
antheridia  can  be  induced 
in  V.  repens  (a  variety  of 
V.  sessilis)  within  4  or  5 
days  by  putting  the  ma- 
terial into  a  2  to  4  per  cent 
cane-sugar  solution  in  ^ 

bright    sunlight.     The    sex  A 

organs  will  not  be  formed        FlG-  39-~ v™che™:    A-  v-  seminata,-   B,  v. 

sessilis;   a,  antheridia;    o,  oogonia. 

in  weak  light  or  in  darkness. 

The  formation  of  zoospores  may  be  induced  in  the  following  way: 
Cultivate  in  a  0 . 1  to  0 . 2  per  cent  Knop's  solution  for  a  week,  then 
bring  the  material  into  tap  water,  and  keep  the  culture  in  the  dark. 
Zoospores  may  appear  within  2  days.  Bright  light  or  a  temperature 
higher  than  15°  C.  will  check  the  production  of  zoospores.  A  2  per 
cent  cane-sugar  solution  kept  in  the  dark  is  also  likely  to  furnish 
zoosporic  material.  If  no  zoospores  are  formed  when  the  solution 
is  kept  in  the  dark,  the  nutrition  has  been  too  weak:  strengthen  the 
nutrient  solution  and  keep  the  culture  in  the  light  for  a  few  days; 
then  put  the  culture  in  the  dark,  and  zoospores  should  appear.  The 
formation  of  zoospores  may  continue  for  a  couple  of  weeks. 

Aplanospores  of  V.  geminata  are  formed  in  nature  when  the  plant 
is  growing  upon  damp  ground.  The  aplanospores  may  also  appear  in 
a  4  per  cent  cane-sugar  solution. 


178  Methods  in  Plant  Histology 

In  fresh  0.5  per  cent  Knop's  solution  in  bright  light,  cultures 
remain  in  the  vegetative  condition,  and  the  result  is  the  same  in  weak 
light  if  the  nutrient  solutions  are  seldom  changed.  Such  cultures 
may  be  kept  indefinitely  by  changing  the  nutrient  solution  whenever 
a  whitish  scum  appears  on  the  surface. 

Fixing  solutions  which  are  successful  with  Spirogyra  may  prove 
ruinous  to  Vaucheria.  In  general,  fixing  solutions  for  Vaucheria 
should  be  very  weak.  Mount  a  few  filaments,  and  with  a  pipette 
add  a  few  drops  of  the  fixing  agent.  Plasmolysis  is  likely  to  take 
place  within  10  to  30  seconds  if  it  is  to  take  place  at  all.  If  plas- 
molysis  takes  place,  weaken  the  fixing  agent.  The  following  formula 
has  given  the  best  results  with  V.  geminata: 

Chromic  acid :  .        1  g. 

Glacial  acetic  acid 8  c.c. 

Water 800  c.c. 

This  is  a  very  weak  solution.  A  loose  mat  as  large  as  one's 
finger  requires  100  c.c.  of  the  reagent.  From  36  to  48  hours  should 
be  allowed  for  fixing.  Wash  24  hours.  Use  the  Venetian  turpentine 
method.  With  Magdala  red  and  anilin  blue  the  oospores  will 
show  various  shades  of  red,  according  to  the  stage  of  development; 
the  filaments  will  show  various  mixtures  of  blue  and  red. 

In  mounting,  use  the  small  scissors  freely.  You  cannot  untangle 
a  mat  of  Vaucheria  so  as  to  give  good  views. 

For  the  development  of  the  oogonium  and  antheridium,  for 
fertilization  and  for  the  structure  and  development  of  the  various 
spores,  thin  sections  are  necessary.  Imbed  in  paraffin.  For  nuclear 
details,  use  iron-haematoxylin ;  f or  cytoplasm,  use  safranin,  gentian- 
violet,  orange. 

Cladophora. — This  genus  is  found  in  both  salt  and  fresh  water. 
The  fresh-water  forms  are  usually  attached  to  sticks  or  stones  in 
quiet  or  running  water.  The  mats  feel  rough  and  crisp  and,  even 
under  a  pocket  lens,  show  the  characteristic  branching  by  which  the 
form  is  easily  recognized.  The  absence  of  a  mucous  coat  makes 
Cladophora  a  convenient  host  for  numerous  parasitic  algae,  among 
which  diatoms  belonging  to  the  genera  Cocconeis  and  Gomphonema 
are  particularly  abundant. 


Chlorophyceae 


179 


For  laboratory  cultures,  select  the  forms  found  in  quiet  water, 
but  for  preparations,  forms  growing  where  the  waves  dash  hard  are 
better,  since  you  can  get  a  fine  display  of  branches  under  a  small 
cover.  Forms  growing  in  still  water  or  in  gently  flowing  water  may 
look  like  unbranched  filaments  under  an  ordinary  cover.  In  fixing, 
use  rather  weak  solutions.  A  chromo-acetic  acid  solution  with  1  g. 
of  chromic  acid  and  4  c.c.  of  glacial  acetic  acid  to  400  c.c.  of  water 
will  usually  produce  no  plasmolysis,  and  will  fix  the  material  in  24 
hours.  After  washing  in  water,  stain  some  of  the 
material  in  iron-haematoxylin  and  put  it  into  the 
10  per  cent  glycerin;  also  put  some  material  in 
without  staining,  reserving  it  for  the  Magdala  red 
and  anilin  blue.  The  iron-haematoxylin  material 
will  give  better  views  of  the  nuclei  and  pyrenoids, 
while  the  other  stain  will  give  better  views  of  the 
other  cell  contents  and  the  cell  walls  (Fig.  40). 

Chara. — Cham  is  found  in  ponds,  lagoons,  and 
ditches.  Once  seen,  it  is  always  readily  recognized 
(Fig.  41).  In  the  ponds  and  lagoons  along  the 
southern  shores  of  Lake  Michigan  it  fruits  so 
abundantly  that  the  whole  pond  shows  an  orange 
color  due  to  the  immense  numbers  of  antheridia. 
In  the  lagoons  of  the  Chicago  parks  Chara  is  so 
abundant  that  it  must  be  dredged  out  every 
summer. 

Chara  is  easily  kept  alive  throughout  the  year 
in  the  laboratory.     A  two-gallon  glass  jar  with  an 
inch  of  pond  dirt,  sand,  and  gravel  at  the  bottom,  and  nearly  filled 
with  tap  water,  is  all  that  is  needed  for  a  successful  culture.     If  the 
jar  is  to  be  covered,  it  should  not  be  more  than  two-thirds  full  of 
water.     Not  more  than  a  dozen  plants  should  be  put  into  such  a  jar. 

A  rather  strong  solution  should  be  used  for  fixing.     The  following 
will  give  good  results: 

Chromic  acid 1  g. 

Glacial  acetic  acid 1  c.c. 

Water...  100  c.c. 


PIG.  40.— Clado- 
phora:  fixed  in 
chromo-acetic  acid 
and  stained  in  iron- 
alum  haematoxylin. 


180 


Methods  in  Plant  Histology 


a.. 


In  about  24  hours  this  not  only  fixes,  but  it  dissolves  the  lime  with 
which  most  species  are  coated. 

For  paraffin  sections  select  the  tip  of  the  plant,  a  piece  about 
half  an  inch  in  length.  Sections  of  this  may  show,  not  only  the  large 
apical  cell,  but  also  various  stages  in  the  development  of  antheridia 
and  oogonia.  For  the  development  of  the  plant  body  from  the 
apical  cell  and  also  for  early  stages  in  the  development  of  oogonia 

and  antheridia,  the 
safranin,  gentian-violet, 
orange  combination  is 
excellent;  for  later 
stages,  especially  in  the 
development  of  the 
antheridia,  iron-haema- 
toxylin  is  much  better. 
The  antheridium  of 
Chara  stains  so  rapidly 
that  the  beginner  uni- 
formly makes  poor  pre- 
parations. In  order  to 
get  good  preparations 
of  the  antheridium,  it 
is  necessary  to  disregard 
other  structures,  which 
will  be  stained  lightly  or  not  at  all  when  the  stan  is  just  right  in 
the  antheridial  filaments. 

If  it  is  desired  to  mount  whole  branches  showing  the  antheridium 
and  oogonium  in  position,  as  in  Fig.  41,  use  the  Venetian  turpentine 
method,  staining  in  Magdala  red  alone,  or  in  Magdala  red  and  anilin 
blue.  Good  mounts  showing  shield,  manubrium,  capitula,  and  fila- 
ments may  be  obtained  by  crushing  an  antheridium  under  a  cover- 
glass.  For  this  it  is  better  to  stain  in  Magdala  red  alone,  since  any 
overstating  is  easily  corrected  by  exposing  the  preparation  to  direct 
sunlight. 


FIG.  41. — Chara:  A,  portion  of  a  branch  showing 
an  antheridium,  a,  and  an  oogonium,  o;  X35;  B,  median 
longitudinal  section  of  an  apical  cell;  drawn  from  a 
preparation  fixed  in  chromo-acetic  acid  and  stained  in 
Delafleld's  haematoxylin;  X225. 


CHAPTER  XV 
PHAEOPHYCEAE.    BROWN  ALGAE 

The  Phaeophyceae,  or  brown  algae,  are  almost  exclusively 
marine.  They  include  a  great  variety  of  forms,  ranging  from  delicate 
filaments  to  coarse,  leathery  plants  a  hundred  feet  in  length.  There 
are  no  unicellular  members. 

For  fixing  marine  algae,  fixing  agents  should  be  made  up  with  sea- 
water,  never  with  fresh  water,  and  the  washing  should  be  done  with 
sea- water;  but  fresh  water  should  be  used  in  making  the  series  of 
alcohols.  When  the  Venetian  turpentine  method  is  to  be  used,  the 
fresh  water  is  used  first  in  making  up  the  10  per  cent  glycerin. 

For  habit  work,  material  may  be  put  into  formalin — about  6  c.c. 
commercial  formalin  to  100  c.c.  of  sea-water — and  kept  there  indefi- 
nitely. If  it  is  desired  to  transport  large  quantities  of  coarse  forms, 
the  material  may  remain  in  this  solution  for  a  week  and  may  then  be 
removed  from  the  liquid  and  packed  in  closed  pails  or  tubs  or  any 
water-tight  containers.  After  reaching  its  destination,  the  material 
should  be  put  into  formalin  again. 

For  material  to  be  mounted  by  the  Venetian  turpentine  method, 
6  to  10  per  cent  formalin  (always  in  sea-water)  is  a  good  fixing  agent. 
Wash  in  sea-water  for  1  hour,  then  in  equal  parts  sea-water  and 
fresh  water  for  ^  hour,  then  in  fresh  water  |  hour.  The  material 
is  now  ready  for  staining  in  aqueous  stains,  or  for  the  10  per 
cent  glycerin,  if  alcoholic  stains  are  to  be  used. 

The  following  formula  by  Flemming  will  also  give  good  results, 
both  for  the  Venetian  turpentine  method  and  for  the  paraffin  method : 

Chromic  acid 1  g- 

Glacial  acetic  acid C,-n: 0.4  c.c. 

Sea-water 400  c.c. 

Fix  24  to  48  hours  and  wash  24  hours  in  running  sea-water.  A 
convenient  washing-box  can  be  made  from  an  ordinary  washtub. 
Bore  a  dozen  f-inch  holes  in  the  bottom;  insert  rubber  tubes  6 

181 


182  Methods  in  Plant  Histology 

inches  long,  and  in  the  end  of  each  tube  place  the  glass  part  of  a 
pipette.  The  tub  may  be  elevated  by  nailing  three  narrow  boards  to 
the  sides  so  as  to  form  a  tripod.  Place  the  bottles  or  cans  of  material 
under  the  pipettes  and  let  sea-water  flow  into  the  tub. 

If  such  chromic  acid  material  is  to  be  used  at  once  for  Venetian 
turpentine  mounts,  follow  the  washing  in  sea-water  by  £  hour's 
washing  in  equal  parts  sea-water  and  fresh  water  (not  necessarily 
running  water)  and  then  £  hour's  washing  in  fresh  water.  The 
material  is  now  ready  for  an  aqueous  stain  or  for  10  per  cent  glycerin. 
If  desirable  to  keep  it  for  future  staining,  put  it  into  5  or  6  per  cent 
formalin  in  fresh  water. 

Material  for  sections  may  be  treated  in  the  same  way,  but  it  is 
often  better  to  add  2  to  10  c.c.  of  1  per  cent  osmic  acid  to  100  c.c. 
of  the  chromic-acid  solution.  The  1  per  cent  osmic  acid  should  be 
made  up  in  distilled  water. 

For  habit  demonstrations  many  of  the  smaller  forms  can  be 
floated  out  and  dried  on  paper.  Ectocarpus,  Desmotrichum,  Dictyota, 
Cutleria,  and  even  small  specimens  of  Laminaria  are  quite  useful 
when  prepared  in  this  way.  Take  a  light  pine  board,  a  little  larger 
than  the  standard  herbarium  sheet,  float  it  in  a  tub  of  water,  place 
on  the  board  the  paper  upon  which  the  material  is  to  be  mounted, 
arrange  the  material  with  a  toothpick  or  the  blunt  end  of  a  needle, 
dipping  all  or  a  part  of  the  board  under  water  whenever  necessary. 
Cover  with  a  piece  of  cheese-cloth,  add  a  blotter  or  two,  as  in  case 
of  flowering  plants,  and  dry  under  gentle  pressure,  changing  the 
blotters  frequently.  The  algae  have  enough  mucilage  to  make  them 
adhere  to  the  paper.  Coarse  forms,  like  Fucus,  may  need  to  be  held 
down  by  strips  of  gummed  paper. 

Sphacelaria. — The  apical  cell  of  Sphacelaria  or  the  nearly  related 
Stypocaulon  affords  an  excellent  study  of  the  structure  of  cytoplasm. 
Flemming's  weaker  solution,  with  the  osmic  acid  even  a  little  weaker 
than  recommended  in  the  formula,  is  good  for  the  apical  cell  and  the 
mitotic  figures,  which  are  quite  conspicuous.  For  these  features 
it  is  a  good  plan  to  break  off  the  tips  so  as  to  have  only  pieces  6  to 
12  mm.  long,  which  will  lie  flat  in  the  paraffin.  The  tips  should  be 
broken  off  after  the  material  has  been  brought  into  xyol.  If 


Phaeophyceae 


183 


whole  tufts  are  imbedded,  the  branches  diverge  enough  to  make 
perfectly  longitudinal  sections  of  the  apical  cells  rather  rare.  Iron- 
haematoxylin  with  a  faint  staining  in  orange  is  a  satisfactory  com- 
bination. 

Ectocarpus. — For  general  morphological  study,  branches  should 
be  mounted  whole  in  Venetian  turpentine.  A  6  to  10  per  cent 
formalin  solution  (in  sea- water,  of  course),  or  the  chromo-acetic  acid 
will  give  good  fixation.  Stain  some  in  iron-haematoxylin  and  some 
in  Magdala  red  and  anilin  blue.  Mount  on  each  slide  some  from 
each  lot.  Unilocular  sporangia  usually  ap- 
pear earlier  than  the  pleurilocular  game- 
tangia.  So  collections  should  be  made  at 
different  seasons.  You  should  have  both 
sporangia  and  gametangia  on  each  slide 
(Fig.  42). 

Desmotrichum. — Forms  as  large  as  Des, 
motrichum  can  be  handled  like  Ectocarpus- 
but  care  must  be  taken  not  to  overstain. 

Laminaria. — In  such  large  forms,  small 
portions  showing  the  structure  and  develop- 
ment of  the  thallus  and  also  the  reproduc- 
tion should  be  cut  out  with  a  razor  and 
then  placed  in  the  fixing  agent.  The 
" sporangia"  of  Laminaria  stain  very  deeply 
and  quickly.  Iron-haematoxylin  is  good, 
but  be  careful  not  to  overstain.  After 
this  stain  is  just  right,  about  3  to  5  minutes 
in  alcoholic  safranin  will  stain  the  muci- 
laginous structures  and  add  to  the  value  of  the  preparation. 

For  habit  study,  small  specimens  up  to  45  cm.  in  length  can  be 
mounted  upon  paper.  They  stick  well  and  seldom  need  to  be  secured 
by  gummed  paper.  Larger  specimens  may  be  allowed  to  dry  and 
may  then  be  stored  away  in  a  box.  When  wanted  for  use,  wet  them 
under  the  tap,  or,  better,  in  salt  water;  after  using,  let  them  dry  and 
return  them  to  the  box.  Specimens  will  stand  four  or  five  such 
soakings  in  fresh  water;  if  a  pint  of  salt  is  added  to  three  or  four 


FIG.  42. — Eclocarpus:  from 
a  preparation  stained  in 
Mayer's  haem-alum;  g,  game- 
tangium;  s,  sporangium. 
X255. 


184 


Methods  in  Plant  Histology 


gallons  of  water,  the  material  may  be  soaked  a  dozen  times  before  it 
passes  its  usefulness.  If  material  has  been  fixed  in  formalin,  it  may 
be  washed  in  sea-water— not  very  thoroughly,  but  enough  to  remove 
the  pungent  odor— and  then  soaked  in  equal  parts  of  glycerin  and 
water.  Use  only  enough  of  the  glycerin  to  make  the  specimens 
flexible,  not  enough  to  make  them  wet  to  handle.  In  this  way, 


FIG.  43. — Cutleria  multifida:  photomicrograph  from  a  preparation  by  Dr.  S.  Yama 
nouchi,  showing  a  sorus  of  oogonia,  each  containing  several  eggs;  thickness,  3  M;  stain 
iron-alum  haematoxylin.  X170. 

material  of  Laminaria,  Macrocystis,  Nereocystis,  Postelsia;  and  other 
large  forms  can  be  kept  in  condition  for  demonstration  and  will  last 
for  years  without  any  attention.  When  not  in  use,  they  should  be 
kept  stored  in  a  box. 

Cutleria. — This  alga  deserves  a  place  in  any  course  in  morphology, 
if  the  course  is  thorough  enough  to  permit  the  study  of  three  members 


Phaeophyceae 


185 


of  the  Phaeophyceae.  These  three  should  be  Ectocarpus  (or 
Pylaiella),  Cutleria,  and  Fucus.  Cutleria  is  not  found  on  the  Ameri- 
can coasts,  but  is  abundant  at  Naples.  The  habits  of  gametophyte 
(known  as  Cutleria)  and  the  sporophyte  (known  as  Aglaozonia)  are 
so  different  that  they  furnish  a  good  illustration  of  alternation  of 
generations.  Beginners  understand  such  an  illustration  more  readily 
than  they  do  an  illustration  like  Dictyota,  with  its  two  generations 
looking  so  nearly  alike.  Cutleria  also  furnishes  a  good  stage  in  the 
evolution  of  sex,  about  midway  between  isogamy  and  the  extreme 
hetrogamy  of  Fucus. 

For  habit  study,  both  generations  should  be  mounted  upon  paper. 
The  gametophyte  (Cutleria) 
sticks  well,  but  the  sporo- 
phyte (Aglaozonia)  will 
need  some  glue  or  gummed 
paper. 

For  paraffin  sections,  fix 
in  chromo-acetic  acid.  Cut 
10  n  thick.  For  mitotic 
figures,  some  osmic  acid 
should  be  added  to  the 
chromo-acetic  acid  and  the 
sections  should  be  much 
thinner,  about  3  to  5  IJL. 
Use  iron-haematoxylin  and 
then  stain  for  3  to  5  minutes  in  alcoholic  safranin  (Figs.  43,  44). 

Fucus. — Material  for  habit  study  may  be  dried,  or  preserved  in 
formalin,  or  mounted  on  paper.  In  the  latter  case,  glue  or  gummed 
paper  will  be  necessary.  Most  satisfactory  of  all  is  to  send  to  Woods 
Hole,  Massachusetts  (George  M.  Gray),  for  living  material.  Fertili- 
zation occurs  at  all  seasons,  but  autumn  is  the  most  favorable.  In 
summer  the  material  dies  before  it  reaches  Chicago,  but  during  the 
rest  of  the  year  a  pailful  will  reach  Chicago,  and  even  as  far  west  as 
the  Mississippi  River,  in  good  condition  for  showing  the  rotation  of 
the  egg  by  the  sperms.  The  eggs  and  sperms  form  slimy  masses, 
the  antheridia  being  orange  red,  and  that  containing  the  eggs  a  dirty 


FIG.  44. — Cutleria  multifida:  photomicrograph 
from  a  preparation  by  Dr.  S.  Yamanouchi,  show- 
ing a  sorus  of  antheridia ;  thickness,  3  n  ;  stain 
iron-alum  haematoxylin.  X170. 


1SI, 


Methods  in  Plant  Histology 


green.  Mix  a  drop  of  the  red  with  a  drop  of  the  green.  The  move- 
ments of  the  egg  can  be  observed,  and  material  for  a  study  of  fertili- 
zation and  later  stages  is  easily  secured.  In  fixing  fertilization  and 
succeeding  stages,  it  is  worth  while  to  use  some  of  the  regular  Flem- 
ming's  weaker  solution,  as  well  as  the  solution  without  the  osmic 
acid. 


FIG.  45. — Fucus  vesiculosus:  A,  small  portion  of  plant  showing  bladders  and  fruiting 
branches;  one-half  natural  size;  B,  transverse  section  of  fruiting  branch  showing  oogonial 
conceptacles ;  X6;  C,  antheridia  and  paraphyses;  from  a  preparation  teased  out  and 
mounted  whole;  X225;  D,  oogonium  showing  five  of  the  eight  eggs;  prepared  as  in  (7. 

For  the  growing  points  and  conceptacles,  small  pieces  should  be 
cut  off  with  a  razor.  If  the  fruiting  tips  be  cut  through  lengthwise 
before  they  are  cut  off,  the  fixing  will  be  more  satisfactory.  For 
sections  of  the  conceptacles  it  is  better  not  to  cut  across  the  whole 
tip,  but  to  cut  off  pieces  of  the  rind  containing  half  a  dozen  concep- 
tacles. Such  pieces  are  more  easily  imbedded  and  cut.  There  is  no 
difficulty  in  cutting  such  pieces  in  paraffin.  Iron-haematoxylin  is  a 


Phaeophyceae 


187 


good  stain.  Safranin  and  gentian-violet  are  also  satisfactory,  but 
care  must  be  taken  not  to  overstain,  since  Fucus  usually  stains  deeply 
and  rapidly. 

For  the  cytologist,  Fucus  might  be  used  as  a  test  object  for  testing 
proficiency  in  technic,  just  as  Pleurosigma  angulatum  is  used  in  test- 
ing an  objective.  The  nuclear  divisions  in  the  antheridium  are 
simultaneous,  and  at  the  sixth  division,  which  is  the  last,  there  are 
32  mitotic  figures,  each  with  32  chromosomes  which  split  so  that  32 
go  to  each  pole.  When  you  can  make  a  preparation  in  which  these 


Fio.  46. — Dictyota  dichotoma:  longitudinal  section  showing  apical  cells;  photo- 
micrograph from  a  preparation  stained  in  iron-alum  haematoxylin.  X 167. 

chromosomes  can  be  counted,  your  technic  is  adequate  for  research 
work  in  cytology  (Fig.  45). 

Dictyota. — Dictyota  deserves  a  place  in  the  series  illustrating  the 
evolution  of  sex,  since  its  large  egg  has  lost  all  motility,  but  the 
difference  in  size  of  egg  and  sperm  is  not  so  extreme  as  in  Fucus. 
It  also  furnishes  an  excellent  example  of  the  development  of  a  thallus 
from  an  apical  cell  (Fig.  46). 

Mount  habit  material  on  paper.  For  sections,  fix  in  chromo- 
acetic  acid.  For  figures,  cut  3  to  5  /*,  but  for  general  views  of  apical 
cell  and  reproductive  phases,  cut  10  ju.  Stain  in  iron-haematoxylin 
and  counter-stain  for  2  or  3  minutes  in  safranin. 


CHAPTER  XVI 
RHODOPHYCEAE.     RED  ALGAE 

The  red  algae  belong  almost  exclusively  to  salt  water,  but  a  few 
genera  are  found  only  in  fresh  water,  usually  in  running  water, 
and  a  few  forms  occur  both  in  salt  and  in  fresh  water.  Nearly  all 
are  small  forms,  and  for  habit  work  can  be  floated  out  and  mounted 
on  paper.  Very  few  will  need  glue  or  gummed  paper. 

For  more  critical  habit  work  and  for  Venetian  turpentine  mounts, 
fix  in  6  to  10  per  cent  formalin  in  sea-water.  Material  keeps 
indefinitely  in  10  per  cent  formalin. 

For  sections,  use  the  chromo-acetic  acid  with  or  without  the  addi- 
tion of  a  little  osmic  acid,  as  recommended  for  the  brown  algae. 
The  same  method  of  fixing  and  washing  should  be  used  as  for  the 
brown  algae,  except  that  in  the  case  of  the  few  fresh-water  forms,  fresh 
water  should  be  used  in  making  the  fixing  agent  and  in  washing  it  out. 
For  Polysiphonia,  and  doubtless  for  many  other  forms,  the  period 
in  the  fixing  agent  should  be  very  much  shortened.  Picric  acid, 
corrosive  sublimate,  and  absolute  alcohol  have  been  tried,  but  the 
results  have  not  been  encouraging. 

Batrachospermum. — This  is  a  green,  fresh-water  member  of  the 
red  algae.  It  is  not  very  uncommon  in  small  streams  (Fig.  47). 
Fix  in  chromo-acetic  acid  (in  fresh  water)  and  use  the  Venetian 
turpentine  method.  Good  preparations  showing  the  nuclei  may  be 
obtained  by  staining  in  Mayer's  haem-alum,  or  Haidenhain's  iron- 
haematoxylin.  After  the  material  is  ready  for  mounting,  tease  out 
a  small  portion,  and  still  further  dissociate  the  filaments  by  tapping 
smartly  on  the  cover. 

Nemalion. — 'Methods  for  preparing  Nemalion  have  been  described 
by  Wolfe.1  Chromo-acetic  acid  proved  to  be  most  satisfactory  for 
fixing.  For  studying  fertilization,  mounts  were  made  as  follows: 
"Young  tips  were  crushed  in  water  under  a  cover-glass  and  on  a  slide 

'Wolfe,  James  J.,  "Cytological  Studies  in  Nemalion,"  Annals  of  Botany,  18:607- 
630.  1904. 

188 


Rhodophyceae 


189 


that  had  previously  been  treated  with  fixative;  the  cover  was  then 
removed,  and  the  water  on  the  slide  allowed  to  evaporate.  The 
gelatinous  nature  of  the  wall  prevents  the  contents  of  the  cell  from 
being  affected  by  this  treatment,  even  when  the  albumen  has  hardened 
sufficiently  to  hold  the  filaments  firmly  in  place."  Stain  in  safranin 
and  gentian-violet,  and  mount  in  balsam. 

Iron-haematoxylin  is  recommended  for  paraffin  sections.  The 
sections  must  be  very  thin, 
5  fj,  or  less.  "Material 
killed  in  2  per  cent  for- 
malin in  sea-water  and 
gradually  transferred  to 
pure  glycerin  kept  its  color 
perfectly." 

It  seems  impossible  to  g 
get  mounts  of  Nemalion  by 
the  Venetian  turpentine 
method.  The  directions 
for  Venetian  turpentine,  in 
the  second  edition  of  this 
book,  were  intended  for 
filamentous  red  algae  in 
general,  but  unfortunately 
the  paragraph  appeared 
under  the  heading,  Nema- 
lion. 

For  mounting  filaments 
without  sectioning,  fix  in 
10  per  cent  formalin,  stain 
in  iron-haematoxylin,  also 
stain  material  in  Magdala 

red  and  anilin  blue,  and  follow  the  glycerin  method,  as  described 
in  chap.  vii.  When  the  material  is  ready  for  mounting,  tease  a  small 
piece  on  the  slide  with  needles,  add  a  round  cover,  and  still  further 
dissociate  the  filaments  by  tapping  on  the  cover, 
size  or  s&me  other  sealing  medium. 


Fid.  47. — Batrachospermum  moniliforme:  from 
a  preparation  stained  in  Mayer's  haem-alum  and 
mounted  in  glycerin;  A,  portion  of  plant  showing 
branches  and  several  cystocarps;  X25;  B,  pro- 
carpic  branch  showing  carpogonium,  p,  and  tricho- 
gyne,  t,  with  a  male  cell,  s,  attached;  X255;  C,  a 
younger  branch  showing  carpogonium  and  tricho- 
gyne;  X255;  D,  branch  with  three  male  cells; 
X255. 


Seal  with  gold 


190 


Methods  in  Plant  Histology 


Polysiphonia.— This  is  a  very  difficult  form  to  handle,  but  Dr. 
Yamanouchi  has  developed  an  adequate  method,  and,  by  following 
it,  anyone  should  be  able  to  get  good  preparations. 

For  mounting  in  glycerin,  glycerin  jelly,  or  in  Venetian  turpentine, 
fix  in  10  per  cent  formalin  and  stain  in  iron-haematoxylin. 

For  sections,  fix  in  Flemming's  weaker  solution,  but  omit  the 
osmic  acid  for  spermatogenesis  and  germination  of  carpospores. 


FIG.  48. — Polysiphonia  fibrillosa:  from  a  preparation  fixed  in  chromo-acetic  acid, 
stained  in  eosin,  and  mounted  in  glycerin;  X255;  A,  an  antheridium;  B,  a  cystocarp. 
with  carpospores;  C,  a  tetrasporic  branch  with  tetraspores. 

The  time  should  be  very  short,  5  to  40  minutes  being  sufficient.  If 
material  is  left  too  long,  it  goes  to  pieces.  Wash  in  a  gentle  stream  of 
sea-water  for  24  hours.  Stain  in  iron-haematoxylin  and  then  for 
2  to  3  minutes  in  safranin  (Figs.  48  and  49). 

With  very  delicate  forms,  like  Callithamnion  and  Griffithsia,  the 
washing  may  be  in  part  or  even  wholly  omitted,  and  the  chromic  acid 
extracted  by  the  lower  alcohols,  the  material  being  kept  in  the  dark. 

Corallina. — Corallina  and  other  forms  whose  surface  is  incrusted 
with  lime  need  special  treatment.  The  following  solution  is  good: 

Chromic  acid 1  g. 

Glacial  acetic  acid 1  c.c. 

Sea-water .  100  c.c*. 


Rhodophyceae 


191 


Fix  24  hours,  changing  the  fixing  agent  2  or  3  times.  Wash  24 
hours  in  sea-water. 

If  carefully  applied,  the  following  is  a  good  method:  Put  the 
material  into  5  per  cent  glacial  acetic  acid  (in  sea-water)  and  watch 


FIG.  49. — Polysiphonia  fibrillosa:  young  cystocarps  showing  carpospores  and  the 
irregular  fusion  cell  beneath;  photomicrograph  from  an  iron-alum  haematoxylin  prepara- 
tion by  Dr.  S.  Yamanouchi.  X125. 

it.  As  soon  as  the  vigorous  effervescence  begins  to  subside,  rinse 
in  sea-water  and  transfer  to  Flemming's  weaker  solution,  and  fix 
24  hours.  Iron-haematoxylin  is  best  for  figures,  but  for  general 
structure  the  safranin,  gentian-violet,  orange  combination  gives 
beautiful  results. 


CHAPTER  XVII 
FUNGI 

In  general,  the  filamentous  fungi  are  treated  like  the  filamentous 
algae,  while  the  fleshy  forms  are  cut  in  paraffin.  Bacteriological 
methods  are  used  in  making  test-tube  and  Petri  dish  cultures. 
Professor  Klebs's  investigations  make  it  easy  to  secure  material  of 
many  forms  in  various  phases  of  their  life  histories. 
PHYCOMYCETES 

Mucor  (Rhizopus). — This  familiar  mold  appears  with  great 
regularity  on  bread.  The  following  is  a  sure  and  rapid  method  for 
obtaining  Mucor:  Place  a  glass  tumbler  in  a  plate  of  water,  put  on 
the  tumbler  a  slice  of  bread  which  has  been  exposed  to  the  air  for 
a  day,  and  cover  with  a  glass  jar.  The  bread  must  not  become 
too  wet. 

To  obtain  a  series  of  stages  in  the  development  of  the  sporangium 
it  is  better  to  use  living  material.  For  class  work,  time  the  cultures 
so  as  to  have  a  plenty  of  sporangia  which  have  not  yet  begun  to 
turn  brown. 

If  permanent  preparations  are  wanted,  they  are  easily  made.  Fix 
for  at  least  24  hours  in  5  to  10  per  cent  formalin;  wash  |  hour 
in  water,  and  then  follow  the  Venetian  turpentine  method.  Eosin, 
Delafield's  haematoxylin,  or  the  Magdala  red  and  anilin  blue  will 
prove  satisfactory. 

The  finer  details  of  the  sporangium  can  be  seen  only  in  thin 
sections.  Mucor  is  the  most  easily  obtained  material  to  illustrate 
the  progressive  cleavage  of  cytoplasm  by  vacuoles.  For  this  pur- 
pose, fix  in  chromo-acetic  acid  (1  g.  chromic  acid  and  2  c.c.  glacial 
acetic  acid  to  200  c.c.  of  water),  with  or  without  the  addition  of 
about  2  c.c.  of  osmic  acid  to  50  c.c.  of  this  solution.  Cut  2  to  5  p. 
in  thickness  and  stain  in  safranin,  gentian- violet,  orange. 

The  zygosporic  stage  in  the  life  history  is  rarely  met  in  nature 
or  in  cultures,  but  when  once  secured  it  may  be  propagated  indefi- 

192 


Fungi 


193 


nitely.  We  have  a  culture  which  has  been  furnishing  illustrative 
material  for  nearly  twenty  years.  Once  in  a  while,  when  a  particu- 
larly good  culture  appears,  lay  aside  some  of  it  to  start  the  next 
culture.  The  best  series  of  stages  generally  appears  between  the 
fourth  and  seventh  days.  Dr.  Blakeslee  shows  why  zygospores  are 
so  infrequent.  The  conjugating  filaments  belong  to  different  strains 
of  mycelia'  which  he  calls 
plus  and  minus  strains,  and 
which,  for  convenience, 
may  be  called  female  and 
male  strains.  The  more 
vigorous  mycelium  is  +', 
and  the  less  vigorous  — . 
When  the  two  strains  come 
together,  zygospores  are 
formed  along  the  line  of 
meeting.  If  +  and  — 
strains  are  started  at  op- 
posite sides  of  a  dish,  they 
will  meet  near  the  middle 
and  form  a  dark  line  of 
zygospores. 

Even  for  elementary 
study,  it  is  worth  while  to 
make  permanent  prepara- 
tions of  the  zygosporic 
stage  (Fig.  50),  Fix  in  5 
to  10  per  cent  formalin. 
Stain  some  in  eosin,  some 
in  Delafield's  haematoxylin,  some  in  Magdala  red  and  anilin  blue, 
and  leave  some  without  any  staining  at  all.  A  slide  with  material 
treated  in  these  four  ways  will  show  all  stages  at  their  best. 

For  sections,  use  the  chromo-acetic  acid  as  indicated  for  sporangia. 
The  nuclei  are  very  small  and  have  never  yet  yielded  much,  although 
many  have  tried  to  study  them.  Professor  Klebs  had  no  success 
in  trying  to  induce  the  zygosporic  condition  in  Mucor. 


FIG.  50.— Rhizopus  nigricans:  various  stages 
in  the  development  of  zygospores  from  a  culture 
on  bread ;  preparation  stained  in  eosin  and  mounted 
in  Venetian  turpentine.  X  80. 


194 


Methods  in  Plant  Histology 


In  the  related  genus,  Sporodinia,  which  is  rather  common  in 
summer  upon  fleshy  fungi,  especially  upon  Boletus  and  its  allies,  the 
zygosporic  condition  is  not  infrequent.  The  very  damp  atmosphere 
and  the  nutrition  necessary  for  the  formation  of  zygospores  may 
be  provided  in  the  laboratory  in  the  following  way:  Put  a  little 
water  in  a  glass  battery  jar  and  place  filter  paper  around  the  inside 
of  the  jar  so  that  it  will  take  up  water  and  thus  keep  the  sides  of  the 
jar  moist.  Place  a  small  beaker  or  dish,  without  any  water  in  it, 
in  the  bottom  of  the  jar,  and  in  the  beaker  place  a  small  piece  of 
bread  dampened  with  the  juice  of  prunes.  Infect  the  bread  with 
spores,  or  use  a  piece  of  bread  upon  which  mycelium  is  already  grow- 
ing. Sections  of  the  root  of  Daucus  carota  may  be  used  instead  of 
the  bread.  Put  a  piece  of  wet  filter  paper  on  a  pane  of  glass  and 
cover  the  jar.  Begin  to  examine  after  24  hours.  The  zygospores 
may  appear  in  4  or  5  days.  A  very  full  account  of  the  methods  by 
which  the  various  phases  of  the  life  history  of  Sporodinia  may  be 
produced  at  will  is  given  by  Klebs  in  the  Jahrbucher  fur  wissen- 
schaftliche  Botanik  32: 1-69,  1898. 

Saprolegnia. — This  is  an  aquatic  mold,  very  common  upon 
insects  and  algae.  Cultures  are  easily  and  quickly  made.  Bring 
in  a  quart  of  water  from  any  stagnant  pond  or  ditch,  and  into  the 
water  throw  a  few  flies.  After  12  to  24  hours  throw  the  water  away, 
rinse  the  flies  in  clean  water,  and  put  them  into  tap  water.  Spo- 
rangia will  probably  appear  within  24  hours.  The  water  must  be 
changed  every  day  to  keep  bacteria  from  ruining  the  culture.  The 
larvae  of  ants  or  small  pieces  of  boiled  white  of  egg  are  better  than 
flies,  if  sections  are  to  be  cut.  Sporangia  may  be  produced  in  the 
greatest  abundance  by  cultivating  the  mycelium  for  several  days 
and  then  transferring  it  to  pure  water  or  to  distilled  water.  As  long 
as  the  nutrient  solution  is  sufficiently  strong  and  fresh,  only  sterile 
mycelium  will  be  produced. 

To  secure  oosporic  material,  mycelium  which  has  been 
highly  nourished  for  several  days  in  a  nutrient  solution  is 
brought  into  a  0 . 1  per  cent  solution  of  leucin,  or  into  a  0 . 05 
to  0 . 1  per  cent  solution  of  haemoglobin.  Begin  to  examine  after 
24  hours. 


Fungi 


195 


Satisfactory  material  for  general  laboratory  purposes  can  be 
secured  as  just  described.  Absolutely  pure  cultures  can  be  secured 
only  by  observing  all  the  precautions  necessary  in  bacteriological 
work. 

Achlya  is  similar  and  equally  good  for  illustrative  purposes. 
It  is  found  on  insects,  fishes,  dead  fish  eggs,  and  on  algae.  The 
zoospores  escape  in  a  mass,  which,  for  a  short  time,  is  held  together  by 
a  transparent  pellicle;  in  Saprolegnia  the  zoospores  swarm  separately. 
In  Saprolegnia,  the  new  sporangia  grow  up  through  the  empty  ones; 
in  Achlya,  the  later  sporangia  arise  on 
lateral  branches  below  the  earlier  ones. 

Fix  in  chromo-acetic  acid.  Stain  some 
in  iron-haematoxylin  and  some  in  Magdala 
red  and  anilin  blue,  using  the  Venetian 
turpentine  method.  For  material  which 
is  to  be  sectioned,  add  a  little  osmic  acid 
to  the  fixing  agent. 

Albugo. — This  fungus  is  quite  common 
on  Cruciferae,  where  the  white  "blisters" 
or  "  white  rust,"  Albugo  Candida,  form 
quite  conspicuous  patches.  Affected  por- 
tions of  leaves  and  stems  should  be  fixed 
in  chromo-acetic  acid  and  cut  in  paraffin. 
Sections  5  ju  or  less  in  thickness  will  be 
found  most  satisfactory.  Stain  in  iron- 
alum  and  counter-stain  lightly  with  orange 
(Fig.  51).  The  oosporic  stage  is  not  so 
easily  recognized,  but  if  the  pods  appear 
distorted  it  will  be  worth  while  to  examine 
them.  The  oosporic  phase  of  Albugo  bliti  is  easily  recognized  on 
Amaranthus,  where  the  oospores  may  be  seen  with  the  naked  eye 
by  holding  the  leaf  up  to  the  light.  The  oospores  usually  occur  in 
more  or  less  circular  patches  upon  the  leaf.  When  they  occur 
among  the  floral  structure,  there  is  often  a  slight  reddish  coloration. 
Unfortunately  for  the  collector,  it  is  very  seldom  that  any  red 
coloration  in  Amaranthus  is  due  to  the  desired  material. 


FIG.  51. — Albugo  Candida 
on  Capsella:  vertical  section 
of  a  blister  on  a  leaf;  prepara- 
tion fixed  in  Flemming's 
weaker  solution  and  stained 
in  safranin,  gentian-violet, 
orange.  X225. 


196 


Methods  in  Plant  Histology 


To  show  the  structure  of  oospheres  and  antheridia,  sections  must 
not  be  thicker  than  5  /z.  Sections  as  thick  as  10  to  15  ju  may  be 
cut  to  show  the  position  of  oogonia  and  antheridia,  although  such 
sections  are  too  thick  to  give  satisfactory  views  of  the  nuclei. 

HEMIASCOMYCETES 

Saccharomyces. — Formerly  it  was  considered  rather  difficult  to 
demonstrate  the  nucleus  of  the  yeast  cell.  With  fresh  growing  yeast 
the  following  method  by  Wager  should  be 
successful:  Fix  in  a  saturated  aqueous 
solution  of  corrosive  sublimate  for  at  least 
12  hours.  Wash  successively  in  water,  30 
per  cent  alcohol,  70  per  cent  alcohol,  and 
methyl  alcohol.  Place  a  few  drops  of 
alcohol  containing  the  cells  on  a  cover, 
and  when  nearly  dry  add  a  drop  of  water. 
After  the  yeast  cells  settle,  drain  off  the 
water  and  allow  the  cells  to  dry  up  com- 
pletely. Place  the  cover,  or  slide,  with  its 
layer  of  cells  in  water  for  a  few  seconds, 
and  then  stain  with  a  mixture  of  fuchsin 
and  methyl  green,  or  fuchsin  and  methyliD 
blue.  Mount  in  glycerin  or  in  balsam. 

ASCOMYCETES 

This  group,  popularly  known  as  the 
"sac  fungi,"  contains  an  immense  number 
of  saprophytic  and  parasitic  forms.  The 
green  mold  on  cheese  and  leather,  the  leaf 
curl  of  peach,  the  black  knot  of  cherry  and 
plum,  and  the  powdery  mildews  are  familiar 
to  everyone.  The  few  objects  selected  will 
enable  the  student  to  experiment,  but  he 
must  not  be  discouraged  if  success  does 

not  crown  the  first  attempt,  for  some  members  of  the  group  present 

real  difficulties. 


FIG.  52. — Peziza  odorata: 
three  asci  and  many  para- 
physes;  fixed  in  corrosive 
sublimate,  stained  in  bulk  in 
alum  carmine,  teased  out, 
and  mounted  in  balsam. 
X245. 


Fungi  197 

Peziza. — The  Pezizas  and  related  forms  are  fleshy,  and  present 
but  little  difficulty  in  fixing,  cutting,  or  staining.  They  are  abundant 
in  moist  places,  on  decaying  wood,  or  on  the  ground.  The  apothecia 
have  the  form  of  little  cups,  which  are  sometimes  black  and  some- 
times flesh-colored,  but  often  orange,  red,  or  green. 

For  general  morphological  work  it  is  better  to  tease  out  fresh 
or  preserved  material.  Such  views  as  that  shown  in  Fig.  52  are  easily 
obtained  in  this  way.  For  permanent  preparations  showing  such 
views,  it  is  better  to  stain  in  bulk  in  alum  carmine  or  in  Delafield's 
haematoxylin,  and  then  tease  out  the  asci  in  glycerin  or  balsam. 
Sections  showing  the  entire  ascus  should  be  10  to  15  fj,  in  thickness. 

For  the  free  nuclear  division  in  the  ascus,  and  also  for  the  develop- 
ment of  the  ascospores,  Flemming's  weaker  solution,  followed  by 
the  safranin,  gentian- violet,  orange  combination  has  given  the  best 
results.  Cyanin  and  erythrosin  are  also  to  be  recommended.  The 
latter  combination  stains  better  when  the  fixing  contains  no  osmic 
acid.  Sections  should  be  3  ju  in  thickness;  if  thicker  than  5  p,  they 
are  likely  to  prove  unsatisfactory  for  any  cytological  study. 

Eurotium. — Eurotium  with  its  conidial  stage,  Aspergillus,  is  a 
very  common  mold  found  on  bread,  cheese,  decayed  and  preserved 
fruit,  etc.  In  the  conidial  stage  it  is  green  and  in  the  ascosporic 
stage,  yellow,  reddish  yellow,  or  reddish  brown.  Aspergillus  is 
almost  sure  to  appear  upon  bread  which  is  kept  moderately  moist, 
because  the  conidia  are  usually  abundant  in  the  atmosphere.  If 
the  bread  be  wet  with  a  10  per  cent  solution  of  cane-sugar  or  with 
grape  juice,  this  stage  appears  sooner  and  in  greater  abundance. 
A  temperature  of  22°  to  30°  C.  is  also  a  favorable  condition. 

The  perithecial  stage  is  not  found  so  frequently,  but  can  generally 
be  secured  by  examining  moldy  preserves.,  However,  if  one  has  the 
mycelium  or  spores,  the  sexual  stage  can  be  induced.  Soak  a  piece 
of  bread  in  a  20  per  cent  solution  of  grape-sugar  in  grape  juice;  upon 
this  sow  the  spores  and  keep  at  a  temperature  of  about  28°  C.  After 
4  or  5  days,  begin  to  examine.  A  40  per  cent  solution  of  cane-sugar 
in  the  juice  of  prunes  is  also  a  good  nutrient  solution. 

For  class  use  or  for  permanent  preparations  it  is  best  to  select 
rather  young  material  which  shows  various  stages  in  development, 


198  Methods  in  Plant  Histology 

from  the  swollen  end  of  the  hypha  to  the  ripe  spore  (Fig.  53).  Per- 
manent preparations  of  the  conidial  stage,  as  shown  in  Fig.  53,  and 
also  of  the  coiled  twisted  filaments  which  initiate  the  ascosporic  stage, 
should  be  made  by  the  Venetian  turpentine  method  or  by  the  glycerin 
method. 

Fix  in  1  per  cent  chromo-acetic  acid  (1  g.  chromic  acid  and  1  c.c. 
acetic  acid  and  100  c.c.  water)  for  24  hours;  wash  in  water  24  hours; 
transfer  to  10  per  cent  glycerin  and  continue  the  Venetian  turpentine 
method. 

Material  may  be  fixed  in  corrosive  sublimate  acetic  acid  (cor- 
rosive sublimate  2  g.,  glacial  acetic  acid  2  c.c.,  and  water  100).  Use 


FIG.  53. — Aspergillus:  from  material  growing  on  a  hectograph  pad ;  fixed  in  chromo- 
acetic  acid,  stained  in  eosin,  mounted  in  glycerin;  A-E,  successive  stages  in  development. 
X375.  All  such  material  is  more  satisfactory  when  mounted  in  Venetian  turpentine. 

it  hot  (85°  C.).  One  minute  is  long  enough.  Wash  in  water  and  add, 
a  few  drops  at  a  time,  the  iodine  solution  used  in  testing  for  starch. 
At  first,  the  brownish  color  caused  by  the  iodine  will  disappear,  but 
after  a  certain  amount  has  been  added  the  brownish  color  will  remain. 
Stain  in  eosin  or  iron-haematoxylin  and  follow  the  Venetian  turpen- 
tine method. 

A  very  rapid  method  for  this  and  for  similar  small  filamentous 
forms  may  be  added.  Forms  as  large  as  Thamnidium  elegans  can 
be  mounted  successfully  by  this  method. 

1.  100  per  cent  alcohol,  2  minutes. 

2.  Eosin  (aqueous),  2  minutes. 

3.  1  per  cent  acetic  acid,  2  to  10  seconds. 

4.  Wash  in  water  5  minutes,  changing  frequently. 

5.  Mount  directly  in  50  per  cent  glycerin  and  seal. 


Fungi  199 

If  the  material  gets  through  the  first  four  stages  without  shrinking 
but  collapses  at  the  fifth,  put  it  into  10  per  cent  glycerin  and  allow 
it  to  thicken  as  usual.  In  either  case,  after  washing  in  water  it  is 
better  to  follow  the  Venetian  turpentine  method. 

All  the  later  perithecial  stages  are  easily  cut  in  paraffin. 

Penicillium. — This  green  mold  is  found  everywhere  upon  de- 
caying fruit,  upon  bread,  and  upon  almost  any  decaying  organic 
substance.  Material  is  even  more  easily  secured  than  in  case  of 
Aspergillus,  and  Penicillium  is  an  easier  type  for  laboratory  study. 
Such  a  satisfactory  study  can  be  made  from  the  living  material  that 
it  is  hardly  worth  while  to  fix  and  stain.  The  very  rapid  method 
described  for  Aspergillus  will  furnish  good  mounts  if  permanent 
preparations  are  desired. 

The  Erysipheae. — The  mildews  are  found  throughout  the  summer 
and  autumn  on  the  leaves  of  various  plants.  Some  of  the  most 
abundant  forms  are  Microsphaera  alni  on  the  common  lilac;  Sphaero- 
theca  castagnei  on  Bidens  frondosa  and  other  species,  on  Erechtites 
hieracifolia,  and  on  Taraxacum  officinale;  Uncinula  necator  on 
Ampelopsis  quinquefolia,  and  U.  salicis  on  Salix  and  Populus; 
Erysiphe  commune  on  Polygonum  aviculare;  and  Erysiphe  cichoria- 
cearum  on  numerous  Compositae  and  Verbenaceae.  For  herbarium 
purposes  they  may  be  preserved  by  simply  drying  the  leaves  under 
light  pressure.  When  needed  for  examination  the  leaf  should  be 
soaked  in  water  for  a  few  minutes,  after  which  the  perithecia  may 
be  scraped  off  and  mounted  in  water.  In  mounting  great  care  must 
be  taken  not  to  break  off  the  appendages.  The  asci  may  be  forced 
out  by  pressing  smartly  on  the  cover  (Fig.  54). 

For  permanent  mounts  of  entire  perithecia  with  appendages,  fix 
in  5  per  cent  formalin  24  hours,  wash  in  water  1  hour,  stain  in  aqueous 
eosin  24  hours,  treat  with  1  per  cent  acetic  acid  1  minute,  wash  thor- 
oughly in  water,  and  then  transfer  to  10  per  cent  glycerin  and  follow 
the  Venetian  turpentine  method.  If  chromic  acid,  corrosive  subli- 
mate, or  alcohol  be  used  for  fixing,  the  appendages  become  brittle  and 
very  easily  break  off.  However,  the  chromo-acetic  mixtures  are 
better  if  it  is  desired  to  make  paraffin  sections  showing  the  develop- 
ing of  the  perithecium  with  its  asci  and  spores.  For  this  purpose 


200 


Methods  in  Plant  Histology 


the  omnipresent  Erysiphe  commune  on  Polygonum  aviculare  is  excep- 
tionally favorable,  because,  after  the  material  has  been  fixed  and  has 
been  brought  into  alcohol,  the  whole  mycelium,  with  the  develop- 
ing perithecia,  may  be  stripped  from  the  leaf  without  the  slightest 
difficulty,  thus  avoiding  the  necessity  of  cutting  the  leaf  in  order  to 
get  the  fungus.  The  stage  in  which  the  perithecia  are  still  white  or 
yellowish  is  the  most  favorable  for  sections.  At  this  stage  the 
material,  when  abundant,  can  be  stripped  off  from  the  leaves  before 


Fia.  54.—  Uncinula  necator  on  Ampelopsis  quinquefolia:  A,  four  asci  containing  asco- 
spores  have  been  forced  out  by  pressing  on  the  cover;  stained  in  fuchsin  and  mounted  in 
balsam;  B,  a  conidiospore;  and  C,  an  appendage  of  Microaphaera  alni,  drawn  from 
living  material.  X192. 

fixing.  Sections  should  not  be  thicker  than  5  /*•  About  3  ju  is  best 
for  free  nuclear  stages  in  the  ascus  and  for  the  development  of  the 
ascospores.  The  safranin,  gentian-violet,  orange  combination  seems 
to  give  the  best  results,  although  cyanin  and  erythrosin  are  quite 
satisfactory  when  the  stains  are  properly  balanced. 

The  Xylariaceae.— Most  of  these  forms,  in  their  mature  con- 
dition, are  black.  In  younger  stages  the  color  is  lighter,  often  show- 


Fungi 


201 


ing  gray,  brick-red,  or  brownish  tints.  Nummularia  is  common  on 
dead  branches  of  beech,  elm,  oak,  locust,  and  other  trees.  It  is  gen- 
erally flat,  orbicular,  or  elliptical  in  form.  Ustilina  is  a  crustaceous 
form,  rather  diffuse  and  irregular  in  shape.  It  is  most  common 
on  the  roots  of  rotten  stumps.  Hypoxylon  is  more  or  less  globose 
in  form,  and  the  color  is  brick-red,  brown,  or  black.  It  is  found  on 
dead  twigs  and  bark  of  various  trees,  especially  beech,  and  is  more 
abundant  in  moist  situations.  Xylaria  (Fig.  55)  is  found  on  decaying 
stumps  and  logs,  and  often  apparently  on  the  ground,  but  really 
growing  on  twigs,  wood, 
and  bark  just  under  the 
surface.  When  mature  it 
is  black  outside  and  white 
or  light-colored  within. 
When  young,  it  is  easily 
cut  in  paraffin;  in  some 
forms  the  ascospores  are 
fully  formed  before  the 
stroma  becomes  hard 
enough  to  occasion  any 
difficulty  in  cutting.  When 
the  stroma  becomes  black, 
many  members  of  the 
Xylariaceae  become  very 
hard  and  brittle,  so  that 
sections  are  likely  to  be 
unsatisfactory.  For  general  morphological  study  it  is  better  to 
break  the  stroma  transversely  and  examine  with  the  naked  eye 
and  with  a  pocket  lens.  The  asci  with  their  spores  can  be  teased 
out  and  mounted  in  water.  For  permanent  preparations,  soak 
the  stroma  for  a  month  in  equal  parts  of  95  per  cent  alcohol 
and  glycerin;  then  cut  sections,  and,  after  leaving  them  in  glycerin 
for  a  day  or  two,  mount  in  glycerin  jelly.  It  is  better  not  to  stain 
the  old  stages  (Fig.  55).  For  illustrative  purposes,  select  forms 
which  can  be  cut  in  paraffin.  The  method  just  given  merely  shows 
that  such  material  can  be  cut. 


FIG.  55. — Xylaria:  A,  transverse  section  of 
young  stroma  showing  perithecia;  X8;  B,  two 
asci  with  ascospores.  X245. 


202 


Methods  in  Plant  Histology 


LICHENS 

The  lichens  are  usually  regarded  as  difficult  forms.  In  younger 
stages  they  occasion  no  trouble,  but  an  old  apothecium  or  a  leathery 
thallus  often  fails  to  cut  well.  By  employing  the  gradual  processes 
already  described  in  chap,  ix,  satisfactory  sections  should  be  ob- 
tained from  thalli  and  mature  apothecia  of  Physcia,  Usnea,  Sticta, 
Collema,  Parmelia,  and  PeUigera. 

Cyanin  and  erythrosin  is  a  very  good  stain  for  lichens.  The 
algae  stain  blue  and  the  filaments  of  the  fungus  take  the  red.  Where 
the  association  of  the  alga  and  the  fungus  is  rather  loose,  as  in 
Dichonema,  more  satisfactory  mounts  can  be  made  by  staining  in 
eosin,  or  haem-alum  and  eosin,  and  then  teasing  slightly  with  needles 
and  mounting  in  glycerin. 

BASIDIOMYCETES 

This  is  an  immense  group,  of  which  the  smuts,  rusts,  mush- 
rooms, toadstools,  puffballs,  and  bracket  fungi  are  the  most  widely 
known  representatives. 

The  Smuts  (Ustilagineae). — The  smuts  are  abundant  on  wheat, 
oats,  corn,  and  various  other  plants. 


FIG.  56.— Puccir 


X12. 


photomicrograph  of  aecidium  stage  on  barberry  leaf. 


The  smuts  may  be  studied  in  the  living  material.  The  following 
method,  described  by  Ellis,  is  worth  remembering:  A  supply  of 
smutted  barley  may  be  obtained  by  sowing  soaked,  skinned  barley 
that  has  been  plentifully  covered  by  Ustilago  spores.  In  such 
material  it  is  easy  to  trace  stages  in  the  development  of  spores. 
Freehand  sections  of  ears  about  12  mm.  long  show  the  mycelium  and 
spore  clusters.  If  smutted  ears  be  removed  and  kept  floating  on  the 


Fungi 


203 


water,  the  spores  continue  to  develop  and  often  germinate.  For 
paraffin  sections  desirable  stages  should  be  fixed  in  Flemming's  fluid 
or  picro-acetic  acid.  Delafield's  haematoxylin,  followed  by  a  very 
light  touch  of  erythrosin  or  acid  fuchsin,  will  give  a  good  stain. 

For  a  study  of  the  germinating  spores  and  conidia,  cultures  may 
be  made  in  beerwort  on  the  slide  or  in  watch  crystals.  Harper's 
method  of  making  preparations  from  such  ma- 
terial is  ingenious  and  will  undoubtedly  prove 
valuable  in  making  mounts  of  various  small  plant 
and  animal  forms.  A  drop  of  the  material  is 
taken  up  with  a  capillary  tube  and  is  then  gently 
blown  out  into  a  drop  of  Flemming's  weaker  solu- 
tion (15  minutes  to  1  hour  was  sufficient  for  the 
fungus  spores).  Cover  a  slide  with  albumen  fix- 
ative, as  if  for  sections.  A  drop  of  the  material, 
without  previous  washing,  is  drawn  up  into  the 
capillary  tube  and  touched  lightly  and  quickly 
to  the  surface  of  the  albumen.  A  series  of  such 
drops,  almost  as  small  as  the  stippled  dots  in  a 
drawing,  may  be  applied  to  the  slide.  The  fixing 
agent  may  now  be  allowed  to  evaporate  some- 
what, but  the  preparation  must  not  be  allowed 
to  dry.  As  the  slide  is  passed  rapidly  through 
the  alcohols,  the  albumen  is  coagulated,  and  the 
preparation  may  be  treated  just  as  if  one  were 
dealing  with  ribbons  of  sections. 

The  Rusts  (Uredineae). — Pucdnia  graminis, 

the  common  rust  of  wheat  and  oats  is  familiar  to  everyone  (Figs. 
56,  57).  The  uredospores,  or  summer  spores,  known  as  the  red  rust, 
and  the  winter  spores,  known  as  the  black  rust,  are  found  in  unfortu- 
nate abundance,  but  the  aecidium  stage  on  the  barberry  is  not  neces- 
sary for  the  vigorous  development  of  rust  in  the  United  States, 
and  is  seldom  found.  Most  teachers  are  obliged  to  depend  upon 
botanical  supply  companies  for  this  material.  There  are,  however, 
various  aecidia  which  are  as  good,  or  even  better,  for  morphological 
study.  The  aecidia  growing  on  Euphorbia  maculata  (spotted  spurge) 


FIG.  57. — Pucdnia 
graminis:  A,  uredo- 
spores on  oats;  B, 
germinating  teleuto- 
spore.  X375. 


204 


Methods  in  Plant  Histology 


are  abundant  and  are  very  easy  to  fix  and  cut.  The  infected  plants 
are  also  very  easily  recognized,  normal  plants  having  the  prostrate 
habit,  while  infected  plants  become  erect  and  the  internodes  become 
greatly  elongated.  Aecidia  growing  on  Arisaema  triphyllum  (Jack- 
in-the-pulpit)  are  also  easy  to  cut.  The  Aecidium  on  Hepatica 
has  large  nuclei  and  affords  particularly 
good  views  of  the  intercalary  cells 
(Fig.  58). 

Flemming's  weaker  solution  is  recom- 
mended for  fixing  and  iron-haematoxylin 
with  a  faint  touch  of  orange  is  a  satis- 
factory stain. 

It  is  rather  difficult  to  get  good  sec- 
tions of  uredospores  and  teleutospores  of 
Puccinia  graminis,  because  the  leaves  of 
wheat  and  oats  are  refractory  objects  to 
cut.  For  illustrative  purposes,  soak  the 
leaves,  scrape  off  the  spores,  and  study 
without  sectioning.  For  sections,  select 
species  growing  on  less  refractory  hosts. 

Everyone  who  studies  the  rusts  should 
attempt  to  germinate  the  uredospores  and 
teleutospores.  For  this  purpose  the 
hanging-drop  culture  may  be  employed, 
as  described  in  the  chapter  on  temporary 
mounts  (chap.  v).  The  uredospores  ger- 
minate readily  all  summer,  but  in  most 
forms  teleutospores  will  germinate  only 
in  the  spring  following  their  maturity. 

However,  the  teleutospores  of  "lepto"  species,  like  Puccinia  xanthii  on 
Xanthium  canadense  (cockle-bur),  will  germinate  as  soon  as  they 
ripen,  and  will  serve  equally  well  for  study.  If  a  particularly  good 
specimen  is  secured,  it  may  be  preserved  by  the  method  previ- 
ously described  for  desmids,  except  that  in  this  case  it  might  be 
worth  while  to  attempt  staining  with  Mayer's  haem-alum,  or  with 
eosin. 


FIG.  58.— Aecidium  on  He- 
patica: fixed  in  chromo-acetic 
acid  with  a  little  osmic  acid,  and 
stained  in  safranin,  gentian- 
violet,  orange;  from  a  prepara- 
tion by  Dr.  Wanda  M.  Pfeiffer. 
X950. 


Fungi 


205 


•  Gymnosporangium,  which  is  rather  common  on  Juniperus  vir- 
giniana  (red  cedar),  forms  its  basidia  in  the  "cedar  apple"  stage. 
Bring  the  yellowish  "cedar  apples"  into  the  laboratory  and  cover 
them  with  a  bell  jar  to  keep  them  moist.  They  may  be  germinating 
when  brought  in;  if  not,  they  will  soon  germinate  in  the  moist 
chamber.  For  mounting  whole,  fix  in 
6  per  cent  formalin  in  water,  stain  in 
iron-haematoxylin,  and  follow  the 
Venetian  turpentine  method.  For 
sections,  fix  in  Flemming's  weaker 
solution. 

The  Fleshy  Fungi.— For  habit 
study,  nothing  is  equal  to  fresh  ma- 
terial; for  second  choice,  buy  canned 
"mushrooms"  (usually  Agaricus  cam- 
pestris)  at  the  grocery;  forms  not 
readily  available  in  field  or  grocery 
may  be  preserved  in  formalin  alcohol 
(6  c.c.  of  formalin  to  100  c.c.  of  50  per 
cent  alcohol).  When  formalin  is  used 
in  water,  the  fungi  become  too  soft. 
Larger  forms  of  the  mushroom,  puff- 
ball,  and  bracket  types  may  be  dried 
in  an  oven.  The  circulation  of  air 
should  be  good  and  the  temperature 
should  be  kept  at  about  50°  C.  After 
drying,  the  fungi  should  be  poisoned. 

For  sections,  Gilson's  fluid  deserves  more  recognition  than  it  has 
received.     It  is  particularly  good  for  soft  forms,  like  Tremella. 

Gilson's  Fluid.— 

95  per  cent  alcohol 42  c.c. 

Water 60  c.c. 

Glacial  acetic  acid 18  c.c. 

Concentrated  nitric  acid 2  c-c- 

Corrosive    sublimate    (saturated    solution    in 

water) n  c-c- 

Fix  about  24  hours  and  wash  in  60  or  70  per  cent  alcohol. 


FIG.  59. — Coprinus:  young  basidia 
with  the  four  nuclei  which,  later,  pass 
into  the  spores ;  fixed  in  chromo-acetic 
acid  and  stained  in  safranin,  gentian- 
violet,  orange.  X780. 


206  Methods  in  Plant  Histology 

Coprinus  micaceus  is  particularly  good  for  a  study  of  gills,  basidia, 
and  the  formation  of  basidiospores,  because  it  is  so  small  that  a  single 
section  may  show  a  fine  series  of  stages.  Gills  which  are  becoming 
brownish  at  the  tip,  but  which  are  still  white  toward  the  top  of  the 
cap,  will  show  a  splendid  series  of  stages.  For  fixing,  cut  out  pieces 
of  the  gills  1  cm.  long  and  3  mm.  thick.  Such  material  fixes  well  in 
Flemming's  weaker  solution.  Cut  paraffin  sections  perpendicular 
to  the  gills.  To  show  the  four  basidiospores,  sections  should  be 
10  to  15  fj.  thick;  to  show  details  of  nuclei,  3  n  is  thick  enough 
(Fig.  59). 

In  Hydnum  and  Polyporus,  cut  out  pieces  about  three  or  four 
spines  or  three  or  four  pores  in  width  and  about  1  cm.  long.  A 
rectangular  piece  which  will  allow  the  transverse  sections  of  the  spines 
or  pores  to  be  about  4  mm.  wide  and  1  cm.  long  cuts  better  than  a 
piece  which  will  give  square  sections. 

In  Boletus,  simply  strip  off  the  hymenium  and  cut  into  pieces 
which  will  give  transverse  sections  of  the  tubes. 

In  Lycoperdon,  Bovista,  Geaster,  and  Scleroderma,  longitudinal 
sections  of  the  entire  fructification  can  be  cut  in  paraffin  as  long  as 
the  fresh  material  is  easily  sliced  with  a  Gillette  blade. 

Young  stages  of  Cyathus,  Crucibulum,  and  Nidularia  cut  easily 
in  paraffin;  somewhat  older  stages  can  be  cut  in  celloidin,  but 
mature  stages  fail  to  cut  by  any  of  our  present  methods. 


CHAPTER  XVIII 
BRYOPHYTES 

The  Bryophytes,  comprising  the  two  groups  of  Liverworts 
(Hepaticae)  and  Mosses  (Musci),  present  a  great  diversity  of  struc- 
ture, some  being  so  delicate  that  good  preparations  are  very  uncer- 
tain, while  others  are  so  hard  that  it  is  difficult  to  get  satisfactory 
sections.  Between  these  extremes,  however,  there  are  many  forms 
which  readily  yield  beautiful  and  instructive  preparations. 

If  but  one  fixing  agent  should  be  suggested  for  the  entire  group, 
it  would  be  chromo-acetic  acid  with  1  g.  chromic  acid  and  2  c.c. 
acetic  acid  to  200  c.c.  of  water.  It  should  be  allowed  to  act  for  about 
24  hours.  For  morphological  study,  excellent  sections  can  be  secured 
from  material  fixed  in  formalin  alcohol,  about  6  c.c.  of  commercial 
formalin  to  100  c.c.  of  70  per  cent  alcohol.  Material  may  be  left 
in  this  solution  until  needed  for  use.  The  convenience  of  this  fixing 
agent  will  hardly  be  appreciated  by  those  who  are  always  within 
reach  of  a  laboratory. 

For  general  study,  the  small,  delicate  forms  may  be  mounted 
whole  in  Venetian  turpentine. 

Instead  of  treating  forms  in  a  taxonomic  sequence,  we  shall 
consider  first  the  gametophyte  structures  under  the  headings  thallus, 
antheridia,  and  archegonia,  and  shall  then  turn  our  attention  to  the 

sporophyte. 

HEPATICAE 

Some  of  the  liverworts  are  floating  aquatics,  but  most  of  them 
grow  on  logs  or  rocks  or  upon  damp  ground.  They  are  found  at  their 
best  in  damp,  shady  places.  Many  of  them  may  be  kept  indefinitely 
in  the  greenhouse.  Ricda,  Marchantia,  Conocephalus,  Asterella,  and 
many  others  vegetate  luxuriously,  and  often  fruit  if  kept  on  moist 
soil  in  a  shady  part  of  the  greenhouse,  and  they  do  fairly  well  in  the 
ordinary  laboratory  if  covered  with  glass  and  protected  from  too 
intense  light.  Ricda  natans  (Ricciocarpus  natans)  is  a  valuable  type 

207 


208 


Methods  in  Plant  Histology 


for  illustrative  purposes.  It  floats  freely  on  the  surface  of  ponds  and 
ditches.  Early  in  the  spring  (during  April  in  the  Chicago  region)  it 
produces  antheridia;  then,  for  a  short  time  (about  the  first  of  May) 
both  antheridia  and  archegonia,  and  still  later  only  archegonia. 
Sporophytes  then  appear  as  black  dots  along  the  grooves.  After  the 
spores  are  shed,  the  thallus  remains  sterile  for  the  rest  of  the  season. 
Marchantia  and  similar  forms  are  not  difficult  to  establish  out  of 
doors.  A  rather  damp,  shady  spot  close  to  the  north  side  of  a  build- 
ing is  best.  Scrapings  from  a  board  which  has  been  nearly  burned 
up  makes  the  best  fertilizer  to  scatter  on  the  soil,  if  one  is  to  cultivate 
Marchantia.  Such  freezing  as  Marchantia  receives  in  the  vicinity 

of  Chicago  does  not  pre- 
vent it  from  appearing 
again  the  next  spring. 
If  it  is  desirable  to  have 
material  throughout  the 
year,  the  out-of-door 
culture  may  be  made 
in  a  box  which  can  be 
brought  into  the  labora- 
tory or  greenhouse  in 
the  winter.  A  box  3 

feet  long,  2  feet  wide,  and  1  foot  deep  will  be  convenient.  It  should 
have  a  glass  cover;  an  old  window  will  do.  There  should  be  about 
six  inches  of  dirt  in  the  box.  A  mixture  of  sand,  loam,  and  charred 
scrapings  will  make  a  good  substratum  for  Marchantia.  If  one  is  to 
raise  liverworts  in  the  laboratory,  it  is  absolutely  necessary  to  note 
carefully  the  conditions  under  which  they  grow  in  the  field. 

The  living  plants  are  very  desirable,  since  they  not  only  furnish 
the  best  possible  material  for  habit  work  and  the  coarser  microscopic 
study,  but  they  also  enable  one  to  secure  complete  series  in  the 
development  of  the  various  organs. 

The  Thallus. — In  many  cases  it  will  not  be  necessary  to  make  a 
special  preparation  for  the  study  of  the  thallus,  since  preparations  of 
antheridia,  archegonia,  or  sporophytes  may  include  good  sections  of 
vegetative  portions.  This  is  particularly  true  of  forms  like  Riccia, 


B 


Fia.  60.— Ptilidium  ciliare:  A,  longitudinal,  and  B, 
transverse  section  of  the  apical  region  of  the  leafy 
gametophyte.  X420. 


Bryoph  ytes — Hepaticae 


209 


where  the  various  organs  are  not  raised  above  the  thallus.  In  forms 
like  Marchantia,  where  the  antheridia,  archegonia,  and  sporophytes 
are  borne  upon  stalked  receptacles,  it  is  better  to  make  separate 
preparations  to  show  the  structure  of  the  mature  thallus.  Sections 
intended  to  show  the  structure  of  the  mature  thallus  should  be  15  n 
to  25  p  in  thickness,  but  sections  to  show  the  growing  point  and 
development  of  the  thallus  should  not  be  thicker  than  10  p.  The 
apical  region  of  the  Jungermanniaceae  (Figs.  60,  61)  affords  an 


Pio.  61. — Pellia  epiphylla:  photomicrograph  of  apex  of  gametophyte,  showing 
apical  cell  and  segments;  safranin,  gentian- violet,  orange.  The  negative  was  made  by 
Dr.  Kohler  at  the  Zeiss  factory  in  Jena,  Germany. 

excellent  opportunity  for  studying  the  development  of  the  plant 
body  from  a  single  apical  cell.  If  mixtures  containing  osmic  acid 
are  used  for  fixing,  there  may  be  difficulty  in  the  staining,  even 
after  using  peroxide  of  hydrogen. 

Chromo-acetic  mixtures,  without  osmic  acid,  are  better  for  the 
apical  region.  Chromo-acetic  acid,  followed  by  Delafield's  haema- 
toxylin,  is  good  for  the  apical  cells  and  developing  regions,  but  a 
light  counter-stain  with  erythrosin  improves  preparations  of  the 
mature  thallus.  In  forms  like  Pellia,  where  even  the  apical  cells 
are  more  or  less  vacuolated,  a  sharp  stain  in  safranin  and  gentian- 
violet  is  quite  satisfactory,  bringing  out  not  only  the  cell  walls,  but 


210  Methods  in  Plant  Histology 

also  the  various  cell  contents  (Fig.  61).  The  chloroplasts  and  leuco- 
plasts  are  well  differentiated  by  this  stain.  After  corrosive  sublimate- 
acetic,  a  vigorous  staining  in  a  mixture  of  acid  fuchsin  and  iodine 
green  often  brings  out  the  walls  very  sharply.  After  corrosive 
sublimate-acetic  the  material  may  be  stained  in  bulk  with  alum 
cochineal  or  alum  carmine,  thus  giving  fairly  good  preparations 
and  saving  considerable  labor. 

Antheridia.— It  is  not  difficult  to  get  good  preparations  showing 
the  development  of  antheridia.     In  forms  like  Conocephalus,  Asterella, 


FIG.  62. — Marchantia  polymorpha:   early  stages  in  the  development  of  antheridia; 
from  an  unpublished  drawing  by  Dr.  W.  J.  G.  Land.     X600. 

Pellia,  etc.,  cut  out  small  portions  of  the  thallus  bearing  the  anther- 
idia. The  piece  should  not  be  more  than  1  cm.  long  and  5  mm.  wide, 
preferably  smaller.  For  the  development  of  the  antheridia  of  M ar- 
chantia,  select  young  antheridiophores  which  still  lie  close  to  the  thallus. 
With  the  antheridiophore,  cut  out  a  small  piece  of  the  thallus,  about 
5  mm.  in  length.  For  general  development,  cut  10  /z,  but  for  details 
of  spermatogenesis,  sections  should  not  be  thicker  than  3  fj.  (Fig.  62). 
If  antherozoids  are  found  escaping,  transfer  them  to  a  small  drop 
of  water  on  a  clean  slide,  invert  the  drop  over  a  1  per  cent  solution 


Bryophytes—Hepaticae 


211 


of  osmic  acid  for  2  or  3  minutes,  allow  the  drop  to  dry  up,  pass  the 
slide  through  the  flame  two  or  three  times,  as  in  mounting  bacteria, 
and  then  stain  sharply  in  acid  f  uchsin.  This  should  show  the  general 
form  of  the  antherozoid,  and  will  usually  bring  out  the  cilia. 

The  Archegonia.— The  methods  for  archegonia  are  practically  the 
same  as  for  antheridia.  Too  much  stress  cannot  be  laid  upon  the 
importance  of  carefully 
selecting  the  material. 
Use  very  small  pieces, 
and,  before  placing  them 
in  the  fixing  agent,  trim 
them  to  such  a  shape 
that  the  position  of 
the  archegonia  will  be 
known  accurately  even 
after  the  pieces  are  im- 
bedded in  paraffin. 

For  stages  like  those 
shown  in  Fig.  63,  A  and 
B,  safranin  with  anilin 

blue   Or  light  green  is  a  FIG.    63.— Marchantia  polymorpha:     A,  three  early 

j    4.   •           j  ij  ,      i  n  stages  in  the  development  of  archegonia — Delafleld's 

gOOd  Stain  and  7  tO  10  /I  haematoxylm;     B,   young  archegonium  showing   two 

is  about  the  right  thick-  neck  canal  ceUs  and  tne  central  cell  before  the  cutting  off 

of  the  ventral  canal  cell — f  uchsin  and  methyl  green ;    C, 

ness,  mature  archegonium  immediately  before  the  fertiliza- 

T?nr  <stno-P«  lilr^  C   in  tion  period — safranin,  gentian-violet,  orange;  D,  young 

3r  Stages  like  C,  in  embryo-Delaneld's  haematoxylin.     X400. 

such  forms  as  Mar- 
chantia, where  the  necks  are  long  and  often  somewhat  curved,  it  is 
better  for  general  purposes  to. use  sections  about  15/i  in  thickness. 
If  it  is  desired  to  obtain  preparations  showing  the  cutting  off  of  the 
ventral  canal  cell,  the  development  of  the  oosphere,  and  the  process  of 
fertilization,  the  sections  should  be  from  3  to  6  /i  in  thickness. 

For  archegonia.  containing  young  embryos,  like  that  shown  in  D, 
Delafield's  haematoxylin  without  any  counter-stain  gives  beautiful 
preparations  when  the  staining  is  well  done.  It  is  easier  for  the 
beginner  to  get  good  preparations  with  the  safranin,  gentian-violet, 
orange  combination. 


212 


Methods  in  Plant  Histology 


In  Riccia  natans  (Ricciocarpus  natans)  the  direction  of  the  axis 
of  the  archegonium  at  every  stage  in  the  development  must  be 
known;  otherwise,  there  will  be  few  good  longitudinal  sections. 

In  forms  like  Porella  and  Scapania,  the  involucre  covering  the 
archegonia  is  likely  to  hold  a  bubble  of  air,  which  will  delay  or  even 
prevent  fixing.  The  best  plan  is  to  cut  off  the  offending  leaf  with 
a  pair  of  slender-pointed  scissors.  Some- 
times the  air  can  be  got  out  with  an  air- 
pump. 

The  Sporophyte. — Sporophytes  in 
early  stages  of  development  often  yield 
good  preparations  without  very  much 
trouble,  but  in  later  stages  they  are 
frequently  difficult  to  cut  on  account  of 
the  secondary  thickening  of  the  capsule 
wall  and  the  stubborn  exine  of  the  ma- 
ture spores.  Great  care  must  be  taken  to 
get  Riccia  natans  into  paraffin  without 
shrinking,  and  the  same  thing  may  be 
said  of  other  forms  which  have  such  loose 
tissue  with  large  air  cavities.  Formerly, 
we  resorted  to  celloidin  for  stages  like  that 
shown  in  Fig.  64.  The  gradual  processes 
already  described  have  obviated  the  diffi- 
culty, so  that  the  student  should  be  able 
to  get  thin  paraffin  sections  as  free  from  distortion  as  were  the 
old  celloidin  sections.  In  Riccia  natans  it  is  even  more  difficult  to 
get  median  longitudinal  sections  of  the  sporophyte  than  of  the  arche- 
gonium. Sections  perpendicular  to  the  groove,  whether  longitudinal 
or  transverse,  are  almost  sure  not  to  give  median  longitudinal  sec- 
tions of  the  sporophyte,  and  these  are  the  sections  the  beginner  is  sure 
to  cut.  Examine  the  material  and  note  very  exactly  the  orientation 
of  the  sporophyte;  then,  for  fixing,  cut  out  sections  about  2  mm. 
thick,  taking  these  sections  in  such,  a  plane  that  paraffin  sections 
parallel  to  the  thick  section  will  give  the  desired  median  longitudinal 
sections  of  the  sporophyte. 


FIG.  64. — Riccia  natans:  young 
sporophyte  inclosed  in  the  arche- 
gonium; spore  mother-cell  stage 
— Delafleld's  haematoxylin.  All 
the  cells  of  the  sporophyte,  ex- 
cept a  single  peripheral  layer 
(dotted  in  the  figure)  produce 
spores.  Celloidin  section  30  ^  in 
thickness.  X104. 


Bryophytes—Hepaticae  213 

In  forms  like  Pellia  and  Aneura,  it  is  desirable  to  show  the  sporo- 
phyte  still  inclosed  in  the  calyptra  (Fig.  65).  For  such  sections,  we 
should  recommend  fixing  in  formalin  alcohol.  Aqueous  fixing  agents 
are  likely  to  cause  trouble  on  account  of  air  bubbles.  For  cytological 
studies,  the  calyptra  must  be  removed  and  a  thin  slab  should  be 
cut  from  opposite  sides  of  the  capsule  to  facilitate  fixing  and  infiltra- 
tion. Chromo-acetic  acid,  with  the 
addition  of  a  little  osmic  acid,  is 
best  for  fixing.  In  Pellia  and  Con- 
ocephalus  the  spores  are  very  large 


FIQ.  65. — Pellia  epiphylla:  photo- 
micrograph of  young  sporophyte  at 
the  spore  mother-cell  stage.  X21. 


Fio.  66. — Pellia  epiphylla:  photomicro- 
graph of  spore  germinating  while  still  re- 
tained within  the  capsule — safranin. 
gentian- violet.  X276. 


and  have  a  rather  thin  wall.  Both  these  genera  show  a  peculiar  intra- 
sporal  development  of  the  gametophyte,i.e.,the  gametophyte develops 
to  a  considerable  extent  before  it  ruptures  the  spore  wall  and  before  it 
is  shed  from  the  capsule  (Fig.  66).  Mitotic  figures  during  the  first 
three  divisions  in  these  spores  are  exceptionally  beautiful  and  are  very 
easy  to  stain  with  the  safranin,  gentian-violet,  orange  combination, 
the  chromosomes  taking  a  very  brilliant  red,  while  the  asters  take 


214 


Methods  in  Plant  Histology 


the  violet.    Achromatic  structures  are  very  prominent  during  these 
three  divisions. 

For  the  older  sporophytes  of  Marchantia,  it  is  better  not  to  cut 
the  whole  receptacle.  Remove  the  radiating  branches.  The 
sporophytes  are  in  radiating  rows,  alternating  with  the  branches.  A 
piece  2  mm.  wide  can  be  cut  so  as  to  include  two  of  the  radiating 
rows,  one  on  each  side  of  the  stalk,  and  such  a  piece  will  include  early 


FIG.  67. — Anthoceros  laetis:  A,  longitudinal  section  of  lower  portion  of  sporophyte 
imbedded  in  the  gametophyte;  X45;  B,  transverse  section  of  lower  portion  of  sporophyte; 
X200;  C,  vegetative  cell  from  lower  portion  of  the  sporophyte;  X560;  D,  spore  mother 
cell  showing  three  of  the  four  chloroplasts  with  numerous  starch  grains;  the  nucleus  in 
the  metaphase  of  the  first  division;  X560. 

stages  in  other  rows.  By  taking  such  care,  you  can  get  median 
longitudinal  sections  of  nearly  all  the  sporophytes.  For  class  work, 
5  to  10  ju  is  a  good  thickness,  but  for  figures,  especially  the  reduction 
mitoses  in  the  spore  mother  cells,  the  sections  should  not  be  thicker 
than  2  or  3  p. 

Among  the  Bryophytes  no  form  affords  a  better  opportunity 
for  studying  the  development  of  spores  than  Anthoceros,  since  a 
single  longitudinal  section  of  the  sporophyte  may  show  all  stages, 
from  earliest  archesporium  to  mature  spores  (Fig.  67).  The  sporo- 


Bryop'hytes — Hepaticae  215 

phyte  is  even  more  difficult  to  orient  than  that  of  Riceia  natans. 
Cut  a  slice  1  or  2  mm.  thick,  so  as  to  orient  the  visible  portion  of 
the  sporophyte,  and  trust  to  luck  for  the  orientation  of  the  foot. 
For  studies  like  A  and  B,  chromo-acetic  material  cut  10  n  thick 
and  stained  in  Delafield's  haematoxylin  is  very  good.  The  starch 
grains  in  the  chloroplasts  take  a  beautiful  violet  color  with  the 
safranin,  gentian-violet,  orange  combination.  With  so  many  stages  in 
a  single  section,  it  will  be  impossible  to  stain  all  of  them  well.  A  stain 
which  will  show  the  mother  cells  and  their  divisions  will  be  too 
deep  for  the  mature  spores,  and  a  stain  which  shows  the  spores 
well  will  be  too  faint  for  the  mother  cells.  It  is  better  to  stain  some 
preparations  for  one  feature  and  others  for  another.  It  is  not  worth 
while  to  steer  a  median  course. 


CHAPTER  XIX 

BRYOPHYTES 

MUSCI 

Material  for  a  study  of  the  mosses  is  much  more  abundant,  and 
a  series  of  stages  in  the  development  of  the  various  organs  is  easily 
secured;  but  it  is  much  more  difficult  to  obtain  good  preparations, 
because  so  many  of  the  structures  are  hard  to  cut.  Chromo-acetic 
acid  is  to  be  recommended  as  the  most  satisfactory  fixing  agent,  but 
where  structures  are  refractory  and  very  likely  to  make  trouble  in 
cutting  it  will  often  be  found  more  satisfactory  to  use  formalin 
alcohol  or  picro-acetic  acid  in  the  70  per  cent  alcohol,  since  material 
fixed  in  these  reagents  does  not  become  as  hard  or  as  brittle  as  that 
fixed  in  any  of  the  chromic-acid  series. 

Protonema. — Protonema  of  some  moss  can  always  be  found  at 
any  season.  Look  for  greenish  patches  resembling  Vaucheria. 
Such  mats  show  the  developing  protonema  and  young  leafy  plants. 
Very  young  mats  of  moss  will  also  show  good  protonema,  but  are 
not  likely  to  show  young  buds.  The  brownish  bulbils,  which  are 
quite  common  in  mosses,  can  be  seen  with  a  good  pocket  lens.  The 
little  Webera,  almost  always  found  on  the  pots  in  the  fernery  or  on 
the  benches  in  greenhouses,  quite  frequently  shows  this  mode  of 
reproduction.  Protonema  is  easily  grown  from  spores. 

Permanent  mounts  are  very  easily  made.  Simply  wash  away 
the  dirt  with  water  and  put  the  material  into  10  per  cent  glycerin, 
and  let  the  glycerin  concentrate.  Mount  in  glycerin  or  glycerin 
jelly  for  permanent  mounts.  Seal  thoroughly.  Such  mounts,  with 
no  fixing  or  staining,  may  retain  the  green  color  for  many  years. 

Antheridia. — It  is  easy  to  find  material  for  a  study  of  antheridia, 
because,  in  so  many  cases,  the  antheridial  plants  can  be  detected  at 
once  without  even  a  pocket  lens.  Funaria,  with  its  bunch  of  anther- 
idia as  large  as  a  pinhead,  is  extremely  common  everywhere.  Spring 
is  the  best  time  to  collect  it,  but  it  is  found  fruiting  in  the  autumn 

216 


Bryophytes — Musci 


217 


and  sometimes  in  summer;  besides,  it  is  easily  kept  in  the  green- 
house, where  it  may  fruit  at  any  time.  Bryum  has  a  still  larger 
cluster  of  antheridia,  which  may  be  seen  at  a  distance  of  several  yards. 
Polytrichum  also  has  a  large  cluster  of  antheridia  surrounded  by 
reddish  leaves,  so  that  the  whole  is  sometimes  called  the  moss 
"flower."  In  making  preparations  of  Polytrichum  these  colored 
leaves  should  be  carefully  removed  after  the  material  has  been  got 
into  70  per  cent  alcohol.  A  single  antheridial  plant  of  Polytrichum 
often  furnishes  a  fairly  complete  series  of  stages  in  the  development 
of  antheridia.  Transverse  sections 
show  not  only  the  antheridia,  but 
also  good  views  of  the  peculiar  leaf 
of  this  genus.  In  all  cases  the  stem 
should  be  cut  off  close  up  to  the 
antheridia,  for  many  of  the  moss 
stems  cut  like  wire. 

Sections  to  show  the  develop- 
ment of  the  antheridium  should  be 
5  to  lO/i  in  thickness.  The  safranin, 
gentian-violet,  orange  is  a  good  combi- 
nation (Fig.  68) .  For  details  of  sper- 
matogenesis,  sections  should  not  be 
thicker  than  3  /z.  Iron-haematoxylin 
is  a  better  stain  for  the  chromatin 
and  blepharoplasts. 

Although  sections  20  to  50  n  in 

thickness  can  be  cut  to  show  topography,  it  is  far  better  to  study 
such  stages  in  the  fresh  material.  When  a  particularly  fine  view  is 
secured  in  this  way,  a  permanent  preparation  may  be  made  by 
putting  the  piece  into  10  per  cent  glycerin,  without  any  fixing  or 
staining,  and  allowing  the  glycerin  to  concentrate.  Then  mount  in 
glycerin  jelly. 

Archegonia. — Since  the  necks  of  the  archegonia  are  usually 
long  and  more  or  less  curved,  it  is  necessary,  for  habit  work,  to  cut 
sections  as  thick  as  20  or  30  /*  in  order  to  get  a  view  of  an  arche- 
gonium  in  a  single  section  (Fig.  68,  A}.  Mayer's  albumen  fixative  is 


B 


Fia.  68. — A,  archegonia  of  Webera 
candicans;  celloidin section  20  M  thick; 
X104;  B,  young  antheridia  of  Poly- 
trichum commune;  X420. 


218  Methods  in  Plant  Histology 

not  likely  to  hold  such  sections  to  the  slide.  Use  Land's  fixative. 
Here,  as  in  case  of  antheridia,  it  is  better  to  use  fresh  material, 
putting  particularly  good  pieces  into  10  per  cent  glycerin  for  glycerin 
jelly  mounts. 

For  the  development  of  the  archegonium,  trim  away  the  leaves 
which  usually  cover  the  cluster.     Fix  in  chromo-acetic  acid  with  a 


Fia.  69. — Funaria  hygrometrica:  A,  apex  of  young  sporophyte  showing  endothecium 
and  amphithecium — chromo-acetic  acid  and  Delafleld's  haematoxylin ;  10  M;  X420; 
B,  C,  and  D,  transverse  sections  of  a  sporophyte  of  the  same  age  as  A,  taken  at  different 
levels;  X255. 

little  osmic  acid  and  cut  5  to  10  /*  thick.  For  a  study  of  the  ventral 
canal  cell  and  fertilization,  sections  should  not  be  thicker  than  3  to  5  ju. 
Sporophyte. — It  is  often  difficult  to  get  good  mounts  of  sporo- 
phytes.  In  the  younger  stages  the  calyptras  are  likely  to  interfere 
with  cutting,  while  in  the  older  stages  the  peristome,  or  hard  wall 
of  the  capsule,  occasions  the  trouble.  If  an  attempt  is  made  to 
remove  the  calyptra  in  young  stages,  like  A  of  Fig.  69,  the  apex 
of  the  sporophyte  usually  comes  with  it.  While  picro-acetic  acid 


Bryophytes — Musti 


219 


material  cuts  more  easily,  chromo-acetic  acid  followed  by  Dela- 
field's  haematoxylin  gives  so  much  sharper  differentiation  in  stages 
like  those  shown  in  Fig.  70  that  it  is  better  to  use  ice  or  Land's  cooler 
and  make  an  effort  to  get  preparations  from  chromic  material. 

Stages  like  that  shown  in  Fig.  70  are  cut  with  comparative  ease, 
for  the  calyptra  is  easily  removed,  and  the  capsule  wall  is  not  yet 


FIG.  70.—Funaria  hygrometrica:  A,  longitudinal  section  of  capsule;  B,  transverse 
section  of  capsule  of  about  the  same  age  as  A— Delafleld's  haematoxylin  and  erythrosin; 
10  n.  The  columella,  archesporium,  outer  spore  case,  two  layers  of  chlorophyll-bearing 
cells,  and  the  beginning  of  the  air  spaces  can  be  distinguished  at  this  stage.  X420. 

hard  enough  to  occasion  any  difficulty.  Safranin,  gentian- violet, 
orange  is  a  good  stain.  The  cell  walls  stain  so  sharply  that  they  are 
not  obscured  by  a  stain  which  will  bring  out  the  cell  contents. 

Later  stages,  after  the  peristome  has  begun  to  differentiate,  are 
likely  to  occasion  difficulty  in  cutting.  Bryum  cuts  as  easily  as 
any  (Fig.  71).  For  the  development  of  the  peristome,  fix  in  formalin 
alcohol  and  stain  in  safranin  and  anilin  blue,  or  in  safranin  and  light 


220 


Methods  in  Plant  Histology 


green.     Safranin  and  Delafield's  haematoxylin  is  also  an  excellent 
stain  for  the  older  stages  in  the  differentiation  of  the  capsule. 

The  mature  sporophytes  of  Sphagnum  (Fig.  72)  are  exceptionally 
hard  to  cut.  It  will  be  worth  while  to  prick  the  capsule  with  a 
needle  when  the  material  is  collected.  This  will  allow  the  fixing 


FIG.  71. — Bryum:  portion  of  nearly 
mature  capsule  showing  operculum,  an- 
nulus,  peristome,  and  three  cells  of  the 
sporogenous  tissue.  X200. 


FIG.  72. — Sphagnum:  longitudinal 
section  of  sporophyte  showing  also 
the  upper  portion  of  the  pseudo- 
podium  and  the  calyptra — Dela- 
fleld's  haematoxylin.  X24. 


agent  to  penetrate  readily,  and  will  also  facilitate  the  infiltration  of 
paraffin  or  celloidin.  The  puncture  causes  only  a  slight  damage, 
and  need  not  reach  the  really  valuable  portion  which  is  to  furnish  the 
median  longitudinal  sections. 

The  younger  stages  in  the  sporophyte  of  Sphagnum,  and  also 
the  antheridia,  archegonia,  and  the  peculiar  development  of  the 
leaves  are  easily  cut  in  paraffin. 


CHAPTER  XX 
PTERIDOPHYTES 

This  group  includes  the  Lycopodiales,  Sphenophyllales,  Psilotales, 
Equisetales,  Ophioglossales,  and  Filicales.  The  Sphenophyllales 
occur  only  as  fossils  and  the  Psilotales  are  confined  to  tropical  and 
subtropical  regions.  The  rest  are  cosmopolitan.  The  Lycopodiales 
are  commonly  called  club  mosses  or  ground  pines,  the  Equisetales 
are  called  horsetail  rushes  or  scouring  rushes,  the  two  common 
genera  of  the  Ophioglossales  are  known  as  the  adder's  tongue 
(Ophioglossum)  and  the  grape  fern  (Botrychiwri) ,  and  the  Filicales 
are  the  common  ferns.  Material  is  abundant,  and  so  easily  recog- 
nized that  anyone  who  pays  a  little  attention  to  collecting  can,  in  a 
single  season,  get  a  fine  supply  for  a  study  of  the  group.  Some 
desirable  forms  may  not  be  present  in  all  localities,  but  these  will  be 
few,  and  can  be  obtained  at  a  reasonable  price  from  those  who  make  a 
business  of  collecting. 

The  technic  for  Sphenophyllales  will  be  found  under  "Special 
Methods"  (chap.  xi).  The  gametophytes  of  Psilotales  are  unknown. 
The  young  sporangia  cut  easily,  but  the  older  stages  should  receive 
great  care  in  dehydrating,  clearing,  and  infiltration.  No  further 
directions  will  be  given  for  these  rather  inaccessible  orders. 

LYCOPODIALES 

Lycopodium. — The  genus  is  evergreen,  and  consequently  some 
stage  in  development  can  be  secured  at  any  season.  In  general,  the 
tropical  species  are  easier  to  cut  than  the  temperate.  Without  any 
regard  to  taxonomic  sequence,  we  shall  consider  the  vegetative 
structure,  the  strobili,  and  the  prothallia. 

Vegetative  structure.— Formalin  alcohol  is  an  excellent  fixing 
agent,  and,  quite  contrary  to  prevalent  notions,  the  staining  capacity 
of  material  seems  to  improve  with  several  months'  immersion. 

The  growing  points  of  stems  and  roots  cut  easily  in  paraffin,  and 
when  the  material  becomes  too  hard  to  cut  in  paraffin  it  can  be  cut 

221 


222 


Methods  in  Plant  Histology 


without  any  imbedding.  It  is  easier  to  get  good  sections  of  L.  lucidu- 
lum  and  L.  inundatum  than  of  drier  species,  like  L.  obscurum  and 
L.  clavatum.  Safranin  and  Delafield's  haematoxylin  is  a  reliable 
stain.  Safranin  with  anilin  blue  or  light  green  is  also  good,  and  the 
light  green  gives  particularly  clear  views  of  the  phloem. 

This  stem,  though  rather  complicated  and  confusing  to  the  begin- 
ner, affords  an  illustration  of  the  exarch  protostele,  the  most  primitive 
type  of  vascular  cylinder  (Fig.  73). 


FIG.  73. — Lycopodium  Billardieri,  a  New  Zealand  species:  photomicrograph  of  trans- 
verse section  of  stem;  fixed  in  formalin  alcohol,  cut  in  paraffin,  and  stained  in  safranin 
and  anilin  blue.  From  a  preparation  by  Dr.  J.  Ben  Hill.  Cramer  contrast  plate; 
16  mm.  objective;  no  ocular  or  AbbS  condenser;  camera  bellows,  1  meter;  yellowish- 
green  filter;  arc  light;  exposure,  5  seconds.  X44. 

The  strobilus. — For  longitudinal  sections,  cut  a  slab  from  each 
side  of  the  strobilus  to  insure  fixing  and  infiltration.  If  a  strobilus, 
or  similar  organ,  is  simply  halved,  both  pieces  are  likely  to  curve. 
Among  north  temperate  species,  Lycopodium  inundatum  is  the  most 
easily  cut.  A  young  strobilus  1  cm.  in  length  may  show  all  stages 
from  the  archesporium  to  the  spore  mother  cell.  Iron-haematoxylin 
is  the  best  stain  for  differentiating  the  archesporial  cells.  The  divi- 


Pteridophytes — Lycopodiaks  223 

sions  in  the  spore  mother  cell  stain  intensely,  so  that  care  must  be 
taken  not  to  overstain. 

The  gametophyte. — In  most  species  the  gametophyte,  or  prothal- 
lium,  is  subterranean,  tuberous,  and  has  no  chlorophyll;  in  other 
species  the  prothallium  is  partly  subterranean  and  partly  aerial,  the 
aerial  portion  being  green  and  bearing  the  archegonia  and  antheridia. 
So  far  as  the  author  is  aware,  no  one  has  ever  found  prothallia  of 
Lycopodium  in  the  United  States,  although  the  prothallia  of  several 
of  our  species,  like  L.  inundatum,  L.  clavatum,  and  L.  annotinum,  are 
well  knowrn  from  European  material.  Nearly  all  the  work  on  Euro- 
pean species  has  been  done  by  Bruchmann,  of  Gotha,  Germany.  No 
one  else  has  ever  found  enough  material  for  any  extended  research. 
He  advises  collectors  to  look,  not  in  dense  patches  of  the  plant,  but 
at  the  edges  of  the  patch.  Look  for  small  plants,  and  if  plants  only 
1  cm.  or  so  in  height  are  found,  then  dig  carefully  for  prothallia. 
With  the  exception  of  L.  inundatum,  forms  which  are  partly  aerial 
have  been  found  only  in  the  tropics. 

It  would  seem  natural  to  get  the  prothallia  by  germinating  the 
spores,  but  here  again  no  one  has  had  any  notable  success,  except 
Bruchmann.  In  some  species,  the  spores  do  not  germinate  for  several 
years,  but  when  the  prothallia  are  once  developed  they  continue  to 
bear  archegonia  and  antheridia  for  several  years.  The  spores  of 
L.  selago  germinate  in  3  to  5  years  after  shedding;  those  of  L.  clavatum 
and  L.  annotinum  in  6  to  7  years.  In  L.  clavatum  and  L.  annotinum 
archegonia  and  antheridia  develop  in  12  to  15  years  after  the  spores 
are  shed.  L.  inundatum  germinates  more  promptly — in  10  days  to 
6  months — but  no  one  has  succeeded  in  keeping  a  culture  up  to  the 
archegonium  stage. 

Botanists  in  Lycopodium  localities  should  look  for  prothallia. 
Since  the  prothallia  of  L.  clavatum  reach  a  length  of  1 . 5  cm.,  it  would 
seem  as  if  they  should  be  found. 

From  material  kindly  furnished  by  Dr.  Bruchmann  it  can  be  said 
that  the  prothallia,  once  secured,  are  easy  to  cut  and  stain. 

Selaginella.— Material  of  Selaginella,  in  all  phases  of  the  life 
history,  is  easy  to  secure,  but  not  so  easy  to  handle  after  it  is  obtained. 
As  many  as  340  species,  mostly  tropical,  have  been  described,  only 


224  Methods  in  Plant  Histology 

three  of  which  are  common  in  the  range  of  Gray's  Manual  Of 
these  three,  Selaginella  apus  is  best  for  sections.  Several  of 
tropical  species  are  common  in  greenhouses  and  they  fruit  abun- 
dantly. 

Vegetative  structure. — Growing  points  and  root-tips  are  easily 
cut  in  paraffin.  In  most  species,  the  older  parts  of  the  stem  are  too 
hard  and  brittle  to  cut  in  paraffin  and  are  too  small  to  cut  well  free- 
hand. Patience  and  a  sharp  knife  seem  to  be  the  only  reliance. 
Some  of  the  tropical  species,  which  have  stems  as  large  as  a  lead 
pencil  and  not  very  hard,  are  best  for  sections.  The  vascular  cylin- 
der is  an  exarch  protostele  and  it  is  exceptionally  easy  to  get  a  sharp 
differential  stain  when  once  the  sections  are  cut. 

The  strobilus. — Very  young  strobili  cut  easily  in  paraffin,  but 
after  the  megaspore  coats  begin  to  harden,  there  are  few  objects 
which  make  more  trouble  than  the  strobili  of  Selaginella.  For  stages 
up  to  the  young  megaspores,  fix  in  chromo-acetic  acid,  with  or  without 
the  addition  of  a  little  osmic  acid;  but  for  later  stages  use  formalin 
alcohol,  and  fasten  the  sections  to  the  slide  with  Land's  fixative. 
Even  the  oldest  stages  can  be  cut  in  paraffin  (Fig.  74). 

The  strobili  of  most  species  are  square  in  transverse  section. 
To  get  longitudinal  sections  showing  the  relations  of  sporangia,  sporo- 
phylls  and  axis,  cut  diagonally,  from  corner  to  corner,  never  paral- 
lel to  the  flat  side.  For  archesporial  cells,  use  iron-haematoxylin; 
for  young  megaspores  and  the  development  of  spore  coats,  use 
safranin,  gentian- violet,  orange;  for  later  stages,  use  safranin  and 
light  green. 

The  gametophytes. — In  some  species,  the  megaspores  and  micro- 
spores  germinate  and  even  develop  up  to  the  egg  and  sperm  stage 
while  still  retained  within  the  sporangia  (Fig.  74).  For  such  stages, 
if  the  strobilus  is  fixed  entire,  use  formalin  alcohol;  if  the  megaspores 
are  removed,  use  chromo-acetic  acid. 

To  make  cultures,  shake  the  spores  out  as  in  case  of  ordinary 
fern  prothallia,  or  scatter  the  whole  strobili  over  the  soil.  The 
female  gametophytes  within  the  old  spore  coats  generally  orient 
themselves  in  the  paraffin,  the  base  of  the  spore  being  down  and  the 
archegonium  end  of  the  gametophyte  being  up. 


Pteridophytes — Lycopodialei 


225 


It  is  hard  to  stain  the  cell  walls  of  the  male  gametophyte. 
blue  is  as  good  a  stain  as  any. 

The  young  embryo,  with  its  two  cotyledons,  its  root,  and  the 
megaspore  still  attached,  makes  an  instructive  preparation  when 
mounted  whole  in  Venetian  turpentine. 


Fia.  74. — Selaginella  apus:  longitudinal  section  of  strobilus  showing  a  microspo- 
rangium  with  germinating  microspores  on  the  left;  on  the  right,  three  of  the  four  mega- 
spores  with  gametophytes  near  the  archegonium  initial  stage;  flxed  in  formalin  alcohol, 
cut  in  paraffin,  and  stained  in  safranin  and  light  green;  from  a  preparation  by  Dr.  W.  J.  G. 
Land.  X80. 

Isoetes. — -This  genus  is  widely  distributed  and  13  of  its  60  species 
occur  within  the  Gray's  Manual  range. 

Vegetative  structure. — The  short,  thick  stem,  even  in  old  plants, 
cuts  easily  in  paraffin.  Fix  in  formalin  alcohol  and  stain  in  safranin 
and  light  green.  Sporelings  with  stems  about  2  mm.  in  diameter  and 
young  plants  with  stems  up  to  5mm.  in  diameter  are  best  for  a 
study  of  the  peculiar  vascular  system  of  this  plant. 


226  Methods  in  Plant  Histology 

Sporangia. — All  the  sporangia  of  the  plant  may  be  said  to  con- 
stitute a  single  strobilus  of  the  Selago  type.  Both  longitudinal  and 
transverse  sections  should  be  cut.  The  stem  is  so  short  that,  in 
a  plant  of  medium  size,  a  longitudinal  section  may  include  the  stem, 
the  sporangium,  and  the  sporophyll,  up  to  the  top  of  the  ligule. 
Such  sections,  10  to  15  p,  or  even  20 /i  in  thickness,  are  best  for 
demonstration.  For  very  detailed  work,  the  older  sporophylls 
should  be  removed  separately,  taking  a  piece  from  the  top  of  the 
stem  to  the  tip  of  the  ligule.  Transverse  sections  through  the  whole 
cluster  of  sporophylls  show  the  arrangement  of  megasporophylls 
and  microsporophylls  and  also  the  relations  of  the  sporangia  to 
sporophylls.  We  have  seen  long,  even  ribbons  through  a  group  of 
old  sporophylls  2.5  cm.  in  diameter  cut  with  a  Gillette  blade  in 
a  Strickler's  holder. 

The  gametophytes. — The  spores  are  shed  in  the  uninucleate  stage, 
and  consequently  it  is  not  so  easy  to  find  the  germination  as  in  the 
case  of  Selaginella.  When  the  large  megasporangium  begins  to 
decay,  let  the  megaspores  dry  naturally.  They  retain  their  power 
of  germination  for  a  year  at  least.  Simply  wet  them  with  tap  water 
and  the  earlier  stages  are  easily  secured,  quite  clean  and  ready  for 
cutting.  There  must  be  soil  in  the  dish  for  later  stages.  Try  a 
similar  method  for  microspores.  Also,  look  at  the  top  of  the  stem 
of  old  plants  for  stages  developing  naturally.  The  cell  walls  of  the 
male  gametophyte,  as  in  the  case  of  Selaginella,  are  rather  hard  to 
differentiate.  Use  anilin  blue  or  light  green. 


CHAPTER  XXI 

PTERIDOPHYTES 
EQUISETALES 

This  order  was  large  and  prominent  in  the  Carboniferous  age, 
but  now  only  a  single  family,  the  Equisetaceae,  survives.  Its  only 
genus,  Equisetum,  contains  24  pieces,  10  of  which  occur  within  the 
Gray's  Manual  range.  Equisetum  is  often  called  the  "scouring 
rush,"  because  the  rough  stems  have  been  used  for  scouring  kettles. 
The  roughness  is  due  to  silica.  Species,  like  E.  hiemak,  which  con- 
tain much  silica,  must  be  treated  with  hydrofluoric  acid  before  the 
older  parts  can  be  cut  in  paraffin. 

Vegetative  Structure. — The  roots  are  very  small,  but  have  large 
cells  and  easily  yield  good  preparations.  If  a  handful  of  Equisetum 
limosum  or  E.  hiemale  growing  in  water  be  pulled  up,  scores  of  root- 
tips  may  be  secured  in  a  few  minutes.  Fix  in  chromo-acetic  acid 
with  a  little  osmic  acid.  In  case  of  such  small  objects  it  is  a  good 
plan  to  add  a  few  drops  of  eosin  to  the  alcohol  during  the  process  of 
dehydrating,  in  order  that  the  material  may  be  seen  more  easily. 
The  slight  staining  does  no  damage,  even  if  more  critical  stains  are 
to  be  used  after  the  sections  are  cut.  Longitudinal  sections  of  the 
roots  may  also  be  obtained  by  cutting  transverse  sections  of  the 
nodes. 

The  growing  points  of  stems  may  be  cut  with  ease  in  paraffin. 
E.  arvense  is  particularly  favorable  on  account  of  the  numerous  apical 
cells  which  may  be  found  in  a  single  preparation. 

The  "fertile"  stem  of  Equisetum  arvense  is  so  free  from  silica 
that  it  can  be  cut  in  paraffin  without  any  difficulty.  The  adult 
vegetative  stem  of  E.  arvense,  and  all  stems  which  contain  so  much 
silica,  must  be  treated  with  hydrofluoric  acid  before  imbedding  in 
paraffin.  However,  nearly  all  of  these  stems  can  be  cut  freehand, 
before  fixing,  without  removing  the  silica.  Fix  freehand  sections 
in  95  per  cent  alcohol.  Material  for  paraffin  sections  should  be 

227 


228 


Methods  in  Plant  Histology 


fixed  in  formalin  alcohol.  Safranin  and  anilin  blue,  with  or  without 
a  little  orange,  is  a  good  combination. 

The  Strobilus. — E.  arvense  affords  the  most  favorable  material 
for  a  study  of  the  development  of  sporangia,  since  the  strobilus  con- 
tains almost  no  silica  and,  even  in  its  latest  stages,  is  easily  cut  in 
paraffin.  In  this  species,  the  young  strobili  are  recognizable  in  July, 
the  sporangia  with  sporogenous  tissue  are  formed  in  August,  and 

the  divisions  in  the  spore  mother 
cell  occur  in  September.  The 
spores  are  not  shed  until  the 
following  April.  If  you  know  a 
patch  of  this  species  which 
"fruits"  every  year,  dig  up 
the  horizontal  underground 
stem  in  July.  The  tip  of  the 
main  axis  is  almost  sure  to  be 
a  strobilus.  Dissect  away  the 
scale  leaves  and  fix  the  strobilus 
in  chromo-acetic  acid  with  a 
little  osmic  acid.  August  and 
September  stages  are  easy  to 
recognize.  If  strobili  are 
brought  into  the  laboratory  in 
December  or  January,  they  shed 
their  spores  within  a  week. 

Strobili  of  other  species,  like 
E.  limosum  and  E.  hiemale,  con- 
tain a  large  amount  of  silica  and, 

consequently,  only  the  younger  stages  cut  well  in  paraffin.  Hydro- 
fluoric acid  damages  the  cell  contents  more  or  less.  In  species  like 
these,  all  stages  in  the  development  are  found  in  a  single  season. 

The  Gametophytes. — The  spores  of  Equisetum  germinate  as  soon 
as  they  are  shed.  They  retain  their  power  of  germination  only  a  day 
or  two.  Shake  the  spores  from  the  strobili  directly  upon  the  soil. 
Sow  spores  in  an  ordinary  flower  pot  in  the  greenhouse,  or  use  a  glass 
dish.  In  the  latter  case,  break  up  pieces  of  flower  pot  for  a  bottom, 


FIG.  75. — Equisetum  arvense:  photomi- 
crograph of  prothallia  with  antheridia. 
X30. 


Pteridophytes — Eguisetales  229 

cover  with  loam,  and  over  this  sprinkle  a  layer  of  sand  about  half 
an  inch  thick.  It  does  no  harm  to  sterilize  everything,  but,  even 
then,  infection  is  sure  to  be  brought  in  with  the  spores.  Con- 
sequently, it  is  a  good  plan  to  wet  the  soil  and  sand  with  water  to 
which  a  little  permanganate  of  potash  has  been  added.  About 
five  or  six  small  crystals  to  a  liter  of  water  is  enough.  Wet  the  soil 
in  this  way,  then  sow  the  spores,  and  cover  with  a  pane  of  glass. 
Antheridia  appear  in  3  to  5  weeks.  Archegonia  appear  later — if 
your  culture  is  not  destroyed  by  blue-green  algae  or  fungi  (Fig.  75). 

For  the  development  of  antheridia,  the  blepharoplast,  and  the 
development  of  the  sperm,  fix  in  Flemming's  weaker  solution  and 
stain  in.  iron-haematoxylin.  The  sperm  of  Equisetum  is  the  largest 
in  Pteridophytes. 

The  prothallia  are  so  small  that  for  morphological  purposes 
it  is  better  to  mount  them  whole.  With  a  knife,  skim  off  a  thin 
layer  of  soil,  just  thick  enough  to  hold  the  prothallia  together.  Fix 
in  formalin  or  in  chromo-acetic  acid  and  stain  some  in  iron- 
haematoxylin  and  some  in  Magdala  red  and  anilin  blue.  Use  the 
Venetian  turpentine  method. 


CHAPTER  XXII 
PTERIDOPHYTES 
OPHIOGLOSSALES 

This  order  contains  only  one  family,  the  Ophioglossaceae,  with 
three  genera,  Ophioglossum,  Botrychium,  and  Helminthostachys.  The 
first  two  species  are  cosmopolitan  but  the  third  is  Australasian. 
Some  regard  this  order  as  merely  a  family,  Ophioglossaceae,  belong- 
ing to  Filicales. 

Botrychium. — Botrychium  is  the  most  available  member  of  the 
order.  While  widely  distributed,  the  individual  plants  are  not 
numerous. 

Vegetative  structures. — The  stem  of  Botrychium  is  erect  and  sub- 
terranean. It  has  an  endarch  siphonostele  with  secondary  growth. 
Cut  away  the  thick,  fleshy  root;  cut  the  stem  into  pieces  about  5  to 
7  mm.  in  length,  fix  in  formalin  alcohol,  and  imbed  in  paraffin. 
Even  the  older  parts  of  old  stems  can  be  cut  in  paraffin  if  you  are 
sufficiently  careful.  Transverse  sections  from  the  base  of  the  bud 
down  to  the  secondary  wood  will  give  a  beautiful  series  in  the  develop- 
ment of  the  stele. 

The  roots,  in  all  stages,  cut  easily  in  paraffin.  The  root-tips 
afford  an  excellent  example  of  development  from  an  apical  cell. 
Fix  in  chromo-acetic  acid.  For  imbedding  in  paraffin,  older  parts 
of  the  root  should  be  cut  into  pieces  not  more  than  5  to  7  mm.  in 
length.  Fix  in  formalin  alcohol.  Transverse  sections  show  a  good 
example  of  exarch  protostele  and  also  of  the  radial  arrangement  of 
xylem  and  phloem  (Fig.  76). 

The  bud  is  a  very  interesting  object.  The  leaf  is  in  its  fourth 
year  when  it  appears  above  ground,  and,  consequently,  the  bud 
contains  young  leaves  of  three  successive  seasons.  Two  of  the  three 
show  a  differentiation  into  sterile  and  fertile  portions. 

Sporangia. — Buds  of  B.  •  virginianum  taken  in  September  or 
October  show  sporangia  with  well-marked  sporogenous  tissue.  For 

230 


Pteridophytes — Ophioglossales 


231 


a  study  of  the  development  of  sporangia,  cut  off  the  fertile  portion 
and  fix  it  separately,  using  Flemming's  weaker  solution  and  staining 
in  iron-haematoxylin.  The  reduction  divisions  in  the  spore  mother 
cell  take  place  after  the  leaves  arrive  above  the  surface.  The  vascu- 
lar system  of  the  sporangium-bearing  portion  and  its  relation  to  the 
rest  of  the  leaf  is  best  shown  by  a  series  of  thick  (about  15  to  20  /*) 
transverse  sections  mounted  on  a  5  X  7-inch  photographic  plate. 


FIG.  76. — Botrychium  mrginianum:  photomicrograph  of  transverse  section  of  root 
showing  exarch  protostele  and  radial  arrangement  of  xylem  and  phloem.  X23. 

They  may  be  covered  by  a  5 X 7-inch  film,  by  a  thin  piece  of  mica, 
or  by  any  thin  piece  of  glass.  Of  course,  cover-glasses  of  this  size 
are  made,  but  are  not  always  available. 

The  gametophyte.—The  gametophyte  of  Botrychium  is  subter- 
ranean and  tuberous.  It  sometimes  reaches  a  length  of  7  to  12  mm. 
and  a  thickness  of  4  to  5  mm.  Usually,  it  is  not  more  than  5  or  6  mm. 
long  and  2  or  3  mm.  thick.  Gametophytes  showing  the  develop- 
ment of  antheridia  and  archegonia  are  not  likely  to  be  more  than  2  or 


232  Methods  in  Plant  Histology 

3  mm.  long  and  1  or  2  mm.  thick.  Near  large  plants,  look  for  small 
sporelings,  not  more  than  1  or  2  cm.  in  height.  Dig  very  carefully 
and  you  may  find  the  gametophytes  attached.  The  soil  should  be 
examined  for  smaller  specimens.  Most  of  the  gametophytes  will  be 
found  at  a  depth  of  1  to  3  cm.  Fix  in  chromo-acetic  acid. 

No  one  has  yet  succeeded  in  raising  the  prothallia  from  the  spores. 
The  prothallia  always  contain  an  endophytic  fungus,  supposed  to  be 
Pythium,  but  even  when  this  is  present  the  spores  do  not  germinate. 

Ophioglossum. — Although  widely  distributed,  our  only  common 
species,  Ophioglossum  vulgatum,  is  so  poorly  represented  in  individuals 
that  it  may  be  regarded  as  a  rather  rare  plant.  The  stem  is  erect 
and  subterranean,  as  in  Botrychium,  but  is  smaller  and  easier  to  cut. 
The  sporangia  are  in  an  unbranched  spike  and  even  the  early  spo- 
rogenous  stages  appear  after  the  leaf  is  above  ground,  so  that  it  is 
comparatively  easy  to  get  material  if  you  know  where  it  grows. 

No  one  has  succeeded  in  finding  prothallia  in  the  United  States. 
Bruchmann,  who  studied  the  Lycopodium  prothallia,  also  found 
and  studied  the  prothallia  of  Ophioglossum  vulgatum.  The  prothal- 
lium  is  circular  in  transverse  section,  about  1  mm.  in  diameter  and 
sometimes  more  than  a  centimeter  in  length.  It  is  subterranean 
and  looks  like  a  small,  irregular  rootlet.  No  one  has  been  able  to 
raise  prothallia  from  spores. 

Prothallia  of  some  of  the  tropical  and  south  temperate  species 
are  easier  to  find,  and  early  stages  have  been  grown  from  spores. 


CHAPTER  XXIII 

PTERIDOPHYTES 
FILICALES 

This  order  includes  the  ferns.  Some  members  are  sure  to  be 
available  in  almost  any  locality  and  all  stages  in  the  life  history  are 
easily  secured. 

Vegetative  Structure. — From  a  technical  standpoint,  the  vegeta- 
tive structures  of  Filicales  present  a  wide  range  of  conditions,  some 
being  so  soft  that  the  greatest  care  must  be  taken  to  get  them  into 
paraffin,  while  others  are  so  hard  that  it  is  almost  impossible  to  cut 
them  at  all. 

The  stem. — Growing  points,  even  of  the  largest  ferns,  can  be  cut 
in  paraffin.  If  the  growing  point  is  covered  with  dense  hairs  or 
ramentum,  either  remove  the  covering  entirely  or,  in  case  of  rather 
fleshy  ramentum,  remove  only  the  scales  which  are  beginning  to  turn 
brownish.  The  white  scales  will  fix  and  cut.  Use  chromo-acetic 
acid.  Unless  mitotic  figures  are  particularly  desirable,  it  is  just  as 
well  not  to  add  any  osmic  acid.  For  illustrating  the  development 
of  the  stem  from  the  apical  cell,  sections  10,  15,  or  even  20  n  are  not 
too  thick. 

Older  portions  of  the  stem,  or  rhizome,  in  most  ferns  are  easily 
cut  while  fresh,  the  sections  being  transferred  to  95  per  cent  alcohol 
after  cutting.  It  is  really  better  to  cut  freehand  the  stems  of  Pteris 
aquilina  and  forms  of  similar  consistency  (Fig.  77).  In  digging  up 
rhizomes,  do  not  merely  dig  down  until  the  rhizome  can  be  grasped 
and  then  pull  it  up,  for  such  material  is  sure  to  show  the  pericycle 
of  the  bundles  torn  away  from  the  parenchyma.  Dig  carefully 
around  the  rhizome  and  then  cut  off  with  a  very  sharp  knife  pieces 
about  two  inches  in  length.  Put  the  fresh  rhizome  into  the  micro- 
tome and  cut  sections  as  thin  as  possible.  Keep  the  knife  wet  with 
water  and  put  the  sections  into  alcohol  as  soon  as  they  are  cut. 
Stain  in  safranin  and  anilin  blue,  safranin  and  light  green,  or  safranin 

233 


234 


Methods  in  Plant  Histology 


and  Delafield's  haematoxylin.  With  any  of  these  combinations,  a 
slight  touch  of  orange  usually  adds  to  the  beauty  of  the  preparation. 
This  particular  stem  affords  a  fine  illustration  of  the  polystele. 
Each  bundle  of  the  polystele  has  the  form  of  a  mesarch  protostele. 
In  Osmunda,  and  in  many  other  ferns  of  similar  habit,  the 
rhizome  is  surrounded  by  the  very  hard  leaf  bases.  Good  sections 


Fia.  77. — Pteris  aquilina:  photomicrograph  of  portion  of  one  of  the  large  bundles 
in  the  rhizome;  cut  freehand  and  stained  in  safranin  and  Delafleld's  haematoxylin. 
X187. 

of  the  central  cylinder  can  be  secured  only  by  dissecting  away  these 
hard  leaf  bases  and  any  hard  portions  of  the  cortex  before  attempting 
to  cut  sections.  A  short  distance  back  of  the  growing  point  will  be 
found  a  region  which  will  show  practically  all  the  structures  of  the 
mature  stem,  which  will  be  easy  to  cut.  Even  in  this  region  the  leaf 
bases  should  be  dissected  away.  From  the  apical  cell  back  to  the 
region  where  the  sclerenchyma  is  beginning  to  turn  brown,  the  ma- 


Pteridophytes—Filicaks  235 

terial  is  easily  cut  in  paraffin.  Older  portions  should  be  cut  freehand. 
Osmunda  affords  an  excellent  illustration  of  the  mesarch  siphonostele. 

For  illustrating  the  amphiphloic  siphonostele,  or  solenostele, 
the  rhizome  of  Adiantum,  the  maiden-hair  fern,  will  furnish  material. 
No  better  material  could  be  found  for  illustrating  the  leaf  gap  and 
leaf  trace. 

The  ferns  of  the  Gray's  Manual  range  afford  no  very  satisfactory 
material  for  illustrating  the  protostele,  although  protosteles  occur 
in  Lygodium  and  Trichomanes.  The  most  satisfactory  material  is 
Gleichenia,  a  very  common  and  very  beautiful  fern  in  tropical  and 
subtropical  regions,  but  almost  never  seen  in  greenhouses  nor  even  in 
botanical  gardens.  Formalin  alcohol  material  is  easily  cut  without 
imbedding  and  is  easy  to  stain. 

The  stems  of  tree  ferns  require  special  treatment.  With  the 
large  leaf  bases  partly  cut  away  with  a  sharp  razor,  transverse  sec- 
tions are  easily  cut  for  a  considerable  distance  below  the  apex. 
Material  fixed  in  formalin  alcohol  cuts  very  well.  If  fresh  material 
is  to  be  cut,  the  softer  portions  should  be  flooded  with  alcohol  after 
each  section.  Farther  down,  there  will  be  a  region  where  sections 
can  be  cut  without  any  flooding,  and  still  farther  down,  it  will  be 
difficult  or  impossible  to  cut  sections  across  the  whole  stem.  Sec- 
tions 1  or  2  cm.  thick,  cut  smooth  on  the  ends,  may  be  kept  in  95  per 
cent  alcohol  or  in  glycerin  in  large  glass  dishes  of  the  Petri  dish 
pattern. 

Roots  are  easy  to  secure  and  easy  to  prepare.  For  mitotic  figures 
and  the  development  of  the  root  from  the  apical  cell,  fix  the  tip  in 
chromo-acetic  acid  with  a  little  osmic  acid.  If  the  development  of 
the  root  is  the  principal  object,  stain  in  safranin  and  light  green,  or 
in  the  safranin,  gentian-violet,  orange  combination;  if  mitotic 
figures  are  to  be  studied,  stain  in  iron-haematoxylin  with  a  very 
light  counter-stain  in  orange. 

Roots  of  tree  ferns  are  sometimes  available  in  greenhouses.  In 
some  species  the  stem  is  covered  by  a  dense  felt  of  small  roots,  some 
of  which  will  be  white  and  soft  at  the  tip.  These  roots  are  likely  to 
have  about  the  diameter  of  onion  root-tips,  and  the  beauty  of  prepara- 
tions made  from  them  could  hardly  be  excelled.  In  the  tropics,  where 


236 


Methods  in  Plant  Histology 


the  plants  are  often  in  the  spray  of  cataracts  and  the  lower  part  of 
the  trunk  is  often  washed  by  mountain  streams,  a  thousand  tips 
might  be  secured  from  a  single  specimen. 

The  roots  of  Angiopteris,  which  become  as  large  as  a  lead  pencil, 
may  be  secured  in  some  greenhouses.    They  cut  easily  after  fixing 


Fio.  78. — Osmunda  cinnamomea:  photomicrograph  of  sporangia  with  spore  mother 
cells  in  various  stages  of  division;  fixed  in  Flemming's  weaker  solution  and  stained  in 
iron-alum  haematoxylin;  from  a  preparation  by  Dr.  S.  Yamanouchi.  X114. 

in  formalin  alcohol  and  furnish  a  fine  example  of  the  exarch  protostele, 
common  to  all  roots. 

The  structure  of  the  leaf  will  appear  in  sections  cut  to  show  the 
sporangia. 

The  Sporangia. — To  illustrate  the  character  of  the  annulus, 
select  sporangia  which  are  just  beginning  to  turn  brown.  Fix  in 
formalin  alcohol  and  dehydrate  as  if  for  paraffin  sections;  after  the 
absolute  alcohol,  transfer  to  10  per  cent  Venetian  turpentine.  Stain- 
ing is  neither  necessary  nor  desirable. 


Pteridophytes—Filicaks  237 

The  various  relations  of  sorus  and  indusium  are  best  illustrated 
by  rather  thick  sections  (10  to  20/z)  of  material  in  which  the  oldest 
sporangia  have  barely  reached  the  spore  stage.  Fix  in  formalin 
alcohol  and  stain  in  safranin  and  anilin  blue. 

For  the  development  of  sporangia,  use  Flemming's  weaker  solu- 
tion. The  sections  should  be  5  to  10  //  in  thickness.  For  the  reduc- 
tion of  chromosomes,  the  sections  should  not  be  thicker  than  3  to  5  p. 
Osmunda  is  particularly  good  for  this  purpose  because  the  number  of 
chromosomes  is  comparatively  small.  The  young  sporangia  of 
Osmunda  dnnamomea  and  0.  Claytoniana  show  the  mother-cell  stage 
in  the  autumn,  but  the  division  into  spores  does  not  occur  until  the 
following  spring,  in  the  vicinity  of  Chicago,  the  mitotic  figures  being 
found  during  the  latter  part  of  April  (Fig.  78).  0.  regalis  does  not 
reach  the  mother-cell  stage  in  the  autumn.  Material  for  mitosis 
should  be  collected  during  the  first  two  weeks  in  May.  Various 
species  of  Pteris  are  common  in  greenhouses  and  are  very  good  for 
development  of  sporangia.  Any  fern  of  the  Aspidium  type  will  yield 
a  good  series,  and  some,  like  Cyrtomium,  may  show  a  fine  series  in  a 
single  sorus.  Marattia,  which  is  likely  to  be  found  in  botanical 
gardens,  will  illustrate  the  "synangium"  type;  Angiopteris  has  a 
sporangium  which  forms  an  easy  transition  to  that  of  the  Cycadales. 

The  Prothallia. — Prothallia  can  usually  be  found  on  the  pots  in 
the  ferneries  of  greenhouses.  Ripe  spores  of  some  fern  or  other  can 
be  obtained  at  any  greenhouse  at  any  time  in  the  year,  and  spores 
of  most  of  our  native  ferns  germinate  well  and  produce  good  pro- 
thallia,  even  if  the  sowing  is  not  made  for  several  months  after  the 
spores  have  been  gathered. 

Fine  prothallia  of  Pteris  aquilina  have  been  grown  two  years 
after  the  spores  were  gathered.  Some,  however,  must  be  sown  at 
once,  or  they  will  not  germinate  at  all.  Spores  which  are  large  and 
contain  enough  chlorophyll  to  make  them  appear  greenish  should 
be  sown  at  once.  The  spores  of  the  common  Osmunda  regalis,  and 
of  the  other  members  of  the  genus,  must  be  sown  as  soon  as  ripe,  or 
they  fail  to  germinate.  The  prothallia  of  0.  regalis,  if  carefully 
covered  with  glass,  may  be  kept  for  a  long  time,  and  they  become 
quite  large.  Prothallia  of  this  fern  in  the  writer's  laboratory  pro- 


238 


Methods  in  Plant  Histology 


duced  ribbon-like  outgrowths  5  mm.  wide  and  more  than  5  cm.  in 
length.  These  prothallia  continued  to  produce  archegonia,  anther- 
idia,  and  ribbon-like  outgrowths  for  more  than  a  year,  when  they 
suddenly  "damped  off."  Lang  watered  prothallia  with  a  weak 
solution  of  permanganate  of  potash,  which  kills  the  fungi  but  does 


B 


Fia.  79. — Pteris  aquilina:  A,  filamentous  stage;  B,  the  apical  cell  has  been  estab- 
lished and  several  segments  have  been  cut  off;  the  figure  shows  the  initial  rhizoid  and 
also  three  rhizoids  coming  from  the  main  body  of  the  prothallium ;  C,  an  older  prothallium 
covered  with  antheridia  in  various  stages  of  development;  from  a  drawing  by  Miss 
M.  E.  Tarrant. 

not  injure  the  prothallia.  He  does  not  state  the  strength  of  the 
solution,  but  four  or  five  crystals  to  a  liter  of  water  seems  to  be 
effective. 

The  prothallia  of  most  ferns  will  grow  for  a  long  time  under  such 
conditions.  Pteris  aquilina  and  many  other  ferns  often  furnish  a 


Pteridophytes — Filicales  239 

good  supply  of  antheridia  in  three  weeks  after  sowing,  and  the 
archegonia  appear  soon  after,  but  it  is  well  to  make  sowings  6  weeks 
before  material  is  needed  for  use.  In  P.  aquilina  and  in  many  others 
if  the  spores  are  sown  too  thickly  only  antheridial  plants  will  be 
obtained  (Fig.  79).  If  prothallia  are  to  produce  archegonia,  they 
must  have  sufficient  room  and  nutrition.  If  there  are  no  green- 
house facilities  and  the  prothallia  must  be  grown  in  the  laboratory, 
it  is  a  good  plan  to  take  a  glass  dish,  10  or  12  inches  in  diameter  and 
about  2  inches  deep,  put  a  layer  of  broken  pieces  of  flower  pots  on  the 
bottom,  cover  this  with  a  layer  of  rich  loam,  and  over  this  sprinkle 
a  layer  of  fine,  clean  sand,  since  sand  is  'much  more  easily  washed 
away  from  the  rhizoids  than  is  the  loam.  The  whole  should  now  be 
thoroughly  wet,  but  not  so  as  to  have  water  standing  on  the  bottom. 
Sow  the  spores  and  cover  with  a  tightly  fitting  pane  of  ground  glass. 
There  should  be  no  need  for  moistening  the  culture  again,  for  pro- 
thallia can  be  kept  fresh  and  vigorous  for  several  months,  or  even  for 
a  year,  without  any  wetting.  When  it  is  desired  to  secure  fertilized 
material,  sprinkle  the  prothallia  with  water,  and  the  young  sporo- 
phytes  will  soon  appear. 

While  there  should  always  be  a  study  from  living  material,  it  is 
worth  while  to  make  permanent  mounts,  even  for  habit  study. 
For  such  study,  the  prothallia  should  be  mounted  whole.  Fix  in 
about  5  per  cent  formalin  and  stain  some  in  iron-haematoxylin  and 
some  in  Magdala  red  and  anilin  blue.  Use  the  Venetian  turpentine 
method.  Each  mount  should  show  the  filamentous  stage,  the 
apical  cell  stage,  and  the  group  of  initials  stage,  and  also  antheridia 
and  archegonia. 

For  paraffin  sections,  it  is  a  good  plan  to  grow  the  prothallia  upon 
Sphagnum,  since  they  can  be  cut  without  removing  them  from  the 
substratum.  Rotten  wood  is  also  a  good  substratum,  since  it  cuts  so 
easily  that  it  is  not  necessary  to  remove  it  from  the  rhizoids. 
Damp  pieces  as  large  as  one's  fist  may  be  placed  under  a  bell  jar, 
where  a  damp  atmosphere  can  be  maintained. 

For  the  development  of  antheridia  and  archegonia,  fix  in  chromo- 
acetic  acid  with  the  addition  of  some  osmic  acid.  If  the  osmic  acid 
is  used  as  strong  as  in  Flemming's  weaker  solution,  allow  it  to  act 


240  Methods  in  Plant  Histology 

for  about  an  hour;  then  transfer  to  the  chromo-acetic  acid  without 
any  osmic.  If  the  gradual  processes  of  dehydrating,  clearing,  and 
infiltrating  have  been  carefully  observed,  about  15  to  20  minutes  in 
the  bath  should  be  sufficient  (Figs.  80,  81,  82). 

For  the  development  of  the  antheridium  and  sperm,  and  especially 
for  the  blepharoplasts  and  their  transformation,  cut  about  3  ju  in 
thickness  and  use  iron-haematoxylin;  for  the  development  of  arche- 
gonia,  cut  at  5  ju  and  stain  in  the  safranin,  gentian-violet,  orange 
combination. 


FIG.  80. — Osmunda  cinnamomea:  photomicrograph  of  vertical  section  from  the 
notch  toward  the  base  of  the  prothallium  showing  four  stages  in  the  development  of 
the  archegonium — chromo-acetic  acid;  safranin,  gentian-violet;  from  a  preparation  by 
Dr.  W.  J.  G.  Land.  X425. 

The  Heterosporous  Filicales. — The  four  genera  Pilularia,  Mar- 
silia,  Salvinia,  and  Azolla  are  aquatic,  the  first  two  growing  rooted 
but  more  or  less  submerged,  and  the  other  two  floating  freely  on 
the  water.  Marsilia  is  the  most  available  and  convenient  labora- 
tory type  of  this  group.  It  is  easily  grown  in  a  pond  or  in  an 
aquarium  in  the  greenhouse.  In  setting  it  out  in  a  pond,  select  a 
place  with  a  gently  sloping  bank,  so  that  part  of  the  material  may  be 
under  water  and  part  may  creep  up  the  bank.  In  the  greenhouse,  a 
rectangular  aquarium  may  be  tilted  to  secure  the  same  conditions. 


Pteridophytes — Filicales 


241 


The  portions  which  are  not  under  water  will  continue  to  fruit  during 
the  summer  and  autumn.  The  whole  sporocarp  cuts  easily  in  paraffin 
during  the  development  of  sporangia,  the  division  of  the  spore 
mother  cells,  and  even  during  the  earlier  stages  in  the  formation  of 
spores.  Except  in  the  case  of  the  youngest  sporocarps,  it  is  better 


PIQ.  82. — Osmunda  cinnamomea: 
photomicrograph  of  a  vertical  sec- 
tion with  a  young  archegonium.  show- 
ing the  neck  canal  cell  with  two 
nuclei,  the  ventral  canal  cell,  the  egg. 
and  the  basal  cell — chromo-acetic 
acid;  safranin.  gentian-violet;  from  a 
preparation  by  Dr.  W.  J.  G.  Land. 
X293. 


FIG.  81. — Osmunda  cinnamomea: 
photomicrograph  of  vertical  section 
of  prothallium  with  an  early  stage  in 
the  development  of  the  archegonium, 
showing  the  basal  cell,  two  neck  cells, 
and,  between  them,  the  cell  which  is 
to  give  rise  to  the  neck  canal  cell,  the  . 
ventral  canal  cell,  and  the  egg — 
chromo-acetic  acid ;  safranin,  gentian- 
violet;  from  a  preparation  by  Dr. 
W.  J.  G.  Land.  X425. 

to  cut  off  a  small  portion  at  the  top  and  at  the  bottom  to  facilitate 
fixing  and  infiltration.  The  mother-cell  stage  and  the  young  spores 
will  be  found  in  sporocarps  which  are  just  beginning  to  turn  brown. 
In  nature,  no  further  nuclear  divisions  take  place  within  the  sporan- 
gium until  the  next  spring,  but  the  wall  of  the  sporocarp  becomes 


242 


Methods  in  Plant  Histology 


extremely  hard.  Sporocarps  for  germinating  should  not  be  collected 
until  there  have  been  one  or  two  sharp  frosts.  The  sporocarps  should 
be  allowed  to  dry  gradually,  after  which  they  may  be  kept  in  a  box 
until  needed  for  use.  They  seem  to  retain  their  power  of  germina- 
tion almost  indefinitely.  Sporocarps  from  poisoned  herbarium 
material  fifty  years  old  have  germinated  readily.  Even  sporocarps 
which  had  been  preserved  in  95  per  cent  alcohol  for  several  years  have 
been  known  to  germinate. 

To  germinate  sporocarps,  cut  away  a  portion  of  the  hard  wall 
along  the  front  edge  and  place  the  sporocarp  in  a  dish  of  water.  The 

gelatinous  ring  with  its  sori 
will  sometimes  come  out  in 
a  few  minutes.  In  less  than 
24  hours  the  microspores, 
starting  from  the  one-cell 
stage,  will  produce  the 
mature  sperms.  The  de- 
velopment of  the  mega- 
spore  is  equally  rapid. 
Embryos  are  abundant  in  2 
or  3  days.  To  secure  a 
series  of  stages  in  the  de- 
velopment of  the  gameto- 
phytes  and  embryo,  it  is 
necessary  to  fix  material  at 
short  intervals. 

For  sections  showing  the  development  of  the  antheridium  and 
sperms,  it  is  better  to  remove  the  megaspores  from  the  sorus,  since 
they  occasion  considerable  difficulty  in  cutting.  Cut  2  to  5  n  in 
thickness  and  stain  in  iron-haematoxylin. 

The  older  megaspores  are  hard  to  cut.  •  It  will  facilitate  infiltra- 
tion and  cutting  if  you  prick  each  megaspore  with  a  sharp  needle 
before  fixing.  Since  the  archegonium  is  at  the  apex  of  the  megaspore, 
the  pricking  need  do  no  damage.  With  good  infiltration  and  Land's 
cooling  device,  smooth  ribbons  can  be  cut,  even  from  such  refractory 
material  (Fig.  83). 


FIG.  83. —  Marsilia  quadri/olia:  upper  portion 
of  megaspore  with  an  archegonium  containing 
a  young  embryo.  X212. 


Pteridophytes — Filicales  243 

The  sperm,  which  in  Marsilia  has  an  unusually  large  number 
of  turns  in  the  spiral,  is  easily  mounted  whole.  When  the  sperms 
have  become  numerous,  put  several  megaspores  upon  a  slide  and. 
heat  gently  until  dry.  Then  wet  the  preparation  in  any  alcohol  and 
stain  sharply  in  acid  fuchsin.  Dehydrate  in  absolute  alcohol  for 
at  least  10  minutes,  clear  in  clove  oil,  and  mount  in  balsam.  Such  a 
preparation  will  often  show  a  score  of  sperms  in  the  gelatinous 
funnel  leading  down  to  the  neck  of  the  archegonium. 

Azolla  is  not  difficult  to  obtain,  and  it  is  easy  to  get  a  series  of 
stages  in  the  development  of  the  micro-  and  megasporangia;  but 
it  is  not  at  all  easy  to  find  the  gametophytes,  since  the  spores  ger- 
minate only  after  they  have  been  set  free  by  the  decay  of  the  plant. 


CHAPTER  XXIV 
SPERMATOPHYTES 

In  variety  of  form  and  in  display  of  individuals,  this  group  sur- 
passes all  others.  We  cannot  hope  to  give  even  approximately 
complete  directions  for  making  preparations,  but  must  be  content 
to  give  a  few  hints  which  may  prove  helpful  in  collecting  material 
and  in  securing  mounts  of  the  more  important  structures.  We 
shall  consider  the  gymnosperms  and  the  angiosperms  separately, 
although  in  many  respects  the  technic  is  the  same  for  both. 

GYMNOSPERMS— CYCADALES 

Material  of  cycads.is  becoming  somewhat  more  available  and 
some  stages  in  the  life  history  can  be  found  in  the  conservatories  of 
city  parks  and  in  botanical  gardens.  In  many  species  the  develop- 
ment of  the  ovule,  and  even  the  development  of  the  female  game- 
tophyte  up  to  the  fertilization  period,  takes  place  quite  naturally 
in  the  greenhouse,  where  pollination  is  not  likely  to  .occur.  The 
development  of  the  staminate  cone  and  pollen  is  perfectly  natural 
under  greenhouse  conditions.  The  vegetative  structures  are  natural 
enough,  but,  with  the  exception  of  leaves  and  small  roots,  are  not 
so  available,  since  material  of  the  stem  would  mean  damage  to  the 
plant. 

The  Vegetative  Structures. — All  the  vegetative  structures  cut 
rather  easily. 

The  stem. — Zamia,1  which  grows  in  various  parts  of  Florida,  is 
the  most  available  material.  Directions  for  handling  the  stem  are 
given  on  p.  125. 

Stems  of  the  larger  cycads  are  not  likely  to  be  obtained,  except 
in  the  field,  and  they  are  confined  to  tropical  and  subtropical  regions. 
They  cut  better  while  fresh;  consequently,  if  one  can  get  material, 
it  is  a  good  plan  to  send  it  to  the  laboratory  and  have  it  cut  before 

1  Material  of  Zamia  pumila  can  be  obtained  at  50  cents  a  plant  (express  collect)  by 
addressing  Mr.  Donald  Murray,  Hawks  Park,  Florida. 

244 


Spermatophytes — Gymnosperms 


245 


fixing.  Even  transverse  sections  are  not  difficult  to  cut  while 
fresh  (Fig.  84).  A  piece  of  cycad  trunk  15  to  30cm.  in  diameter 
and  20  cm.  in  length  will  survive  a  journey  of  six  weeks  or  even 
two  months,  if  care  be  taken  to  coat  the  exposed  ends  with  a  mixture 
of  melted  paraffin  and  moth  balls,  using  three  or  four  moth  balls  as 
large  as  marbles  to  half  a  kilo  of  paraffin.  If  material  is  to  be  fixed 
before  cutting,  use  6  to  10  per  cent  formalin  in  water. 


FIG.  84.— Dioon  spinulosum:  photomicrograph  of  transverse  section  of  wood  of 
Dioon  spinulosum,  cut  from  fresh  material.  X105. 

The  course  of  the  vascular  bundles,  as  they  pass  to  the  cones,  is 
quite  peculiar.  Instructive  preparations  may  be  made  by  cutting 
longitudinal  sections,  about  3  mm.  thick,  through  the  apex  of  the 
stem  and,  without  staining,  clearing  thoroughly  and  mounting  in 
balsam.  In  this  way  we  have  mounted  sections  5  cm.  long,  15  cm. 
wide,  and  3  mm.  thick. 

The  root. — Small  roots,  up  to  a  centimeter  in  diameter,  are  easily 
cut  freehand.  The  tender  root-tips  and  also  the  peculiar  "root- 
tubercles"  should  be  fixed  in  chromo-acetic  acid  and  imbedded  in 
paraffin. 


246 


Methods  in  Plant  Histology 


The  leaves. — The  young  tender  leaves  should  be  fixed  in  formalin 
alcohol  and  imbedded  in  paraffin.  The  adult  leaves  are  rigid  and 
cut  well  freehand.  Stain  sharply  in  safranin,  extract  the  stain  until 
it  almost  entirely  disappears  from  the  cellulose  walls,  then  stain  in 
light  green. 

Spermatogenesis.— Except  in  the  earliest  stages,  the  staminate 
cones  are  too  large  to  be  cut  whole.  The  individual  sporophylls, 


FIQ.  85.— Ceratozamia  mexicana:  A,  pollen  grain  which  has  been  in  a  sugar  solution 
for  two  days;  X876;  B,  nucellus  with  numerous  pollen  tubes;  X17;  C,  basal  end  of 
pollen  tube  showing  the  persistent  prothallial  cell;  outside  it  the  stalk  cell;  and,  above, 
the  two  sperms  still  inclosed  in  the  sperm  mother  cells;  X156. 

with  their  sporangia,  cut  easily  up  to  the  formation  of  microspores; 
then  the  sporangium  wall  hardens  rapidly  and  cutting  becomes 
difficult.  Up  to  the  young  microspore  stage,  fix  in  chromo-acetic 
acid;  fix  later  stages  in  formalin  alcohol.  Transverse  sections  are 
more  instructive  and  are  much  more  easily  cut,  since  the  peripheral 
end  of  the  sporophyll  can  be  cut  only  in  younger  stages. 


Spermatophytes — Gymnosperms  247 

In  all  the  genera  of  cycads,  the  microspore  germinates  while 
still  within  the  sporangium,  the  pollen  grain  at  the  time  of  shedding 
consisting  of  a  prothallial  cell,  a  generative  cell,  and  a  tube  cell.  For 
preparations  at  the  shedding  stage,  shake  the  cone  over  a  piece  of 
paper  and  pour  the  pollen  into  the  fixing  agent.  Use  formalin  alco- 
hol, unless  you  have  so  much  pollen  that  you  can  lose  most  of  it 
and  still  have  enough  left. 

The  pollen  germinates  readily.  Shake  some  of  it  into  a  10  per 
cent  solution  of  cane-sugar  in  water.  In  two  or  three  days  the  pollen 
tubes  will  appear  and  in  a  week  or  two  may  grow  to  several  times  the 
length  of  the  grain;  but,  as  far  as  we  know,  the  generative  cell  has 
not  divided  in  sugar  solutions  (Fig.  85,  A). 

The  development  of  the  pollen  tube  and  its  structures  must  be 
studied  in  sections  of  the  nucellus.  As  soon  as  the  integument  is 
removed  the  nucellus  is  exposed  and  the  position  of  the  pollen  tubes 
is  easily  determined,  since  the  haustorial  portions  of  the  tubes  form 
brownish  lines  radiating  from  the  nucellar  beak.  Having  learned 
the  location  of  the  pollen  tubes,  it  is  better  not  to  remove  the  integu- 
ment, but  to  remove  the  female  gametophyte;  then  cut  from  the 
under  side  of  the  nucellus  against  the  hard,  stony  layer  of  the  integu- 
ment so  as  to  remove  a  small  piece  of  the  nucellus  5  to  7  mm.  square, 
according  to  the  species.  Fix  in  Flemming's  weaker  solution  with 
the  osmic  acid  somewhat  weaker  than  the  formula  indicates  (Fig. 
85,  B  and  C,  and  Fig.  86). 

The  pollen  tubes,  with  their  sperms,  make  instructive  prepara- 
tions when  mounted  whole.  Fix  the  nucellus,  with  its  pollen  tubes, 
as  if  for  paraffin  sections.  About  6  per  cent  formalin  in  water  has 
proved  successful.  Wash  in  water  for  half  an  hour  and  stain  in 
aqueous  safranin,  1  or  2  hours.  Extract  the  stain  until  it  is  satis- 
factory, and  then  transfer  to  10  per  cent  glycerin  and  follow  the 
Venetian  turpentine  method.  When  the  turpentine  becomes  thick 
enough  for  mounting,  tease  the  pollen  tubes  from  the  nucellus  and 
mount  with  pieces  of  cover-glass  under  the  cover  to  prevent  crushing. 

A  skilful  technician,  with  steady  hands,  can  tease  out  the  pollen 
tubes  before  fixing.  It  is  easier  to  judge  the  stain  if  the  pollen  tubes 
are  removed  in  this  way. 


248 


Methods  in  Plant  Histology 


Oogenesis.— The  ovules  of  the  Cycads  and  Ginkgo  are  very  large, 
and,  when  mature,  thin  sections  cannot  be  cut  by  any  method  yet 
discovered.  In  younger  stages  it  is  not  difficult  to  get  good  sections 


FIG.  86. — Ceratozamia  mexicana:  photomicrograph  of  a  section  of  a  mature  sperm 
showing  the  large  nucleus  (very  black  in  the  photo),  the  cytoplasmic  sheath  with  sections 
of  the  ciliated  band  and  cilia;  fixed  in  Flemming's  weaker  solution  and  stained  in  iron- 
alum  haematoxylin.  X473. 


of  the  entire  ovule.  Slabs  should  be  cut  from  two  sides  of  the  ovule 
to  facilitate  fixing  and  infiltration.  During  free  nuclear  stages  in  the 
endosperm,  and  even  during  earlier  stages  in  the  formation  of  walls, 
care  must  be  taken  that  the  slabs  may  not  cut  into  the  endosperm, 


Spermatophytes — Gymnosperms  249 

which  is  so  turgid  that  distortion  would  be  sure  to  result.  Even  after 
the  ovule  approaches  its  full  size,  it  can  be  cut  entire,  until  the  stony 
layer  begins  to  harden.  Paraffin  sections  of  the  entire  ovule,  cut 
15  to  20  n  thick,  and  stained  rather  lightly  in  safranin,  gentian- 
violet,  orange,  make  very  instructive  preparations.  When  the  fresh 
ovule  can  no  longer  be  cut  easily  with  a  razor,  it  is  not  worth  while 
to  try  to  cut  it  in  paraffin.  Interesting  preparations  may  be  made 
by  cutting  from  the  median  longitudinal  portion  of  the  ovule  a  slab 
about  5  mm.  thick.  The  slab  should  be  fixed,  washed,  dehydrated, 
and  cleared  in  zylol.  It  should  then  be  kept  in  a  flat-sided  bottle. 


FIG.  87. — Zamia  floridana:  photomicrograph  of  a  small  portion  of  the  proembryo 
showing  simultaneous  free  nuclear  division— safranin,  gentian- violet,  orange;  Cramer 
contrast  plate;  4mm.  objective;  ocular  X4;  yellowish-green  filter;  Camera  bellows, 
50cm.;  exposure,  6  seconds.  X413. 

Such  a  preparation  shows  the  integument,  micropyle,  nucellus  with 
its  beak,  pollen  tubes,  the  stony  and  fleshy  layers,  general  course 
of  vascular  bundles,  and  the  female  gametophyte  with  its  arche- 
gonia. 

For  thin  sections  of  the  archegonia,  a  cubical  piece  with  an 
edge  of  6  or  8  mm.  should  be  cut  from  the  top  of  the  endosperm  with 
a  veiy  sharp,  thin  blade.  The  slightest  pressure  upon  the  archegonia 
will  ruin  the  preparations. 

Sporophyte.— During  the  period  of  simultaneous  free  nuclear 
division,  which  follows  the  fertilization  of  the  egg,  the  mitotic 
figures  are  quite  striking  and  are  easily  stained  (Fig.  87). 


250  Methods  in  Plant  Histology 

After  the  embryos  begin  to  grow  down  into  the  endosperm, 
oblong  pieces  containing  the  embryos  should  be  cut  out. 

After  the  cotyledons  appear,  useful  preparations  may  be  made  by 
dissecting  out  the  entire  embryos,  which  may  be  fixed  in  chromo- 
acetic  acid,  washed,  stained  in  eosin  or  in  Delafield's  haematoxylin, 
placed  in  10  per  cent  glycerin,  and  mounted  by  the  Venetian  turpen- 
tine method.  Since  the  suspensors  become  long  and  irregular,  each 
embryo  should  be  placed  in  a  separate  dish,  lest  the  suspensors 
become  entangled  and  broken. 

After  the  stony  layer  becomes  hard,  it  is  better  to  use  a  small 
fret  saw  for  opening  the  ovule.  Before  the  embryo  has  pushed  down 
into  the  endosperm,  the  ovule  should  be  sawed  in  two  transversely. 
The  endosperm  and  nucellus  can  then  be  picked  out  and  treated  as 
desired.  After  the  tip  of  the  embryo  reaches  the  middle  of  the  endo- 
sperm, the  ovule  should  be  sawed  open  longitudinally. 

GYMNOSPERMS— GINKGOALES 

From  the  standpoint  of  technic,  the  Ginkgoales,  now  represented 
only  by  Ginkgo  biloba,  are  less  difficult  than  the  Cycadales,  but  the 
difficulties  are  somewhat  similar. 

The  Vegetative  Structures. — The  adult  stem  is  harder  to  cut  than 
Pinus,  but  good  sections  should  be  secured  by  boiling  in  water  and 
soaking  for  a  few  days  in  glycerin.  Transverse  sections  of  the  "spur" 
shoots  are  easily  cut.  They  have  a  comparatively  large  pith  and 
narrow  zone  of  wood,  thus  contrasting  sharply  with  a  long  shoot  of 
the  same  diameter,  which  has  a  small  pith  and  wide  zone  of  wood. 

Spermatogenesis. — The  entire  staminate  cone,  even  at  the  time  of 
shedding  pollen,  can  be  cut  in  paraffin.  For  the  latest  stages,  how- 
ever, it  is  better  to  remove  the  sporophylls  and  cut  them  separately, 
since  the  sections  must  not  be  thicker  than  5  ;u,  if  they  are  to  show 
the  internal  structures  of  the  pollen  grain. 

The  young  staminate  cones  become  recognizable  in  June;  by 
September,  they  have  nearly  or  quite  reached  the  spore  mother-cell 
stage,  but  the  division  of  the  spore  mother  cell  does  not  take  place  un- 
til the  following  April.  In  these  early  stages  the  bud  scales  should  be 
carefully  dissected  away  before  fixing.  Pollen  is  shed  early  in  May. 


Spermatophytes — Gymnosperms  251 

Pollen  tubes  and  their  structures  must  be  studied  in  sections  of 
the  nucellus.  Fertilization,  in  the  Chicago  region,  occurs  about  the 
middle  of  September. 

Oogenesis. — Young  ovules  about  0.25mm.  in  length  are  found 
about  the  middle  of  April;  the  megaspore  mother  cell  and  its  divi- 
sion into  four  megaspores  are  found  about  the  first  of  May;  the 
free  nuclear  stage  in  the  development  of  the  female  gametophyte 
extends  from  the  first  week  in  May  to  the  first  week  in  July;  during 
July,  walls  appear;  then  the  archegonium  initials  and  the  growth  of 
the  archegonium,  the  ventral  canal  cell  being  cut  off  the  second  week 
in  September;  fertilization,  free  nuclear  division  in  the  sporophyte, 
and  the  beginning  of  walls  may  all  be  found  before  the  end  of  Sep- 
tember; cotyledon  stages  belong  to  October,  and  when  the  seeds 
fall  in  November  the  embryo  extends  throughout  nearly  the  entire 
length  of  the  endosperm.  This  is  the  winter  resting  stage,  but, 
planted  in  the  greenhouse,  the  seeds  germinate  without  any  resting 
period,  as  in  the  case  of  cycads. 

For  all  stages  in  oogenesis  and  development  of  the  embryo,  use 
chromo-acetic  acid.  The  free  nuclear  stages  in  both  gametophyte 
and  sporophyte  are  almost  sure  to  plasmolyze.  The  addition  of  a 
little  osmic  acid  may  minimize  the  danger.  Chromo-acetic  acid, 
with  the  chromic  rather  weak  and  the  acetic  rather  strong,  may  fix 
without  plasmolyzing,  and  thus  give  better  views  of  the  general 
structure;  but  material  fixed  in  any  reagent  containing  a  large  per- 
centage of  acetic  acid  is  not  likely  to  be  satisfactory  for  a  study  of 
chromatin. 

For  sections  of  the  entire  ovule,  use  safranin,  gentian-violet, 
orange;  for  free  nuclear  stages  in  both  gametophyte  and  sporophyte, 
use  iron-haematoxylin  with  a  touch  of  orange;  for  the  megaspore 
membrane,  safranin  seems  to  be  the  best  stain. 

GYMNOSPERMS— CONIFERALES 

Since  Pinus  is  an  available  laboratory  type,  we  shall  describe 
methods  for  demonstrating  various  phases  in  the  life  history  of  this 
genus,  hoping  that  the  directions  will  enable  the  student  to  experi- 
ment intelligently  with  similar  forms.  The  dates  are  for  Pinus 


252  Methods  in  Plant  Histology 

Laritio  in  the  vicinity  of  Chicago,  but  dates  will  be  different  for  differ- 
ent species  and  even  for  the  same  species  in  different  regions;  P. 
Laricio,  at  Chicago,  sheds  pollen  about  the  middle  of  June,  but  P. 
maritima  at  Auckland,  New  Zealand,  sheds  its  pollen  about  the 
first  of  October.  After  a  year's  collecting  in  any  region,  there  should 
be  no  difficulty,  since  the  dates  do  not  vary  much  from  year  to  year. 

The  Vegetative  Structures. — The  stem,  root,  and  leaf  will  be 
treated  separately. 

The  stem. — The  vascular  cylinder  is  an  endarch  siphonostele,  a 
type  which,  with  few  exceptions,  is  found  throughout  the  living 
gymnosperms. 

The  young  stem  in  its  first  year's  growth  is  green  and  soft  and  is 
easily  cut  in  paraffin.  The  best  time  to  collect  material  is  soon  after 
the  young  shoot  has  emerged  from  the  bud  scales  in  the  spring.  With 
a  thin  safety-razor  blade,  cut  the  stem  transversely  into  pieces  about 
5mm.  in  length;  fix  in  formalin  alcohol,  imbed  in  paraffin,  and 
stain  in  safranin  and  light  green.  Longitudinal  sections  of  the  buds 
in  winter  or  early  spring  condition  are  instructive  for  comparison 
with  longitudinal  sections  of  the  ovulate  cone.  Trim  away  most  of 
the  bud  scales  and  cut  a  slab  from  opposite  sides,  leaving  a  piece 
2  or  3  mm.  thick  to  be  imbedded.  The  bud,  and  also  selected  pieces 
of  the  young  stem,  will  show  the  structure  of  the  young  leaf.  Later 
in  the  season,  even  the  first  year's  shoot  should  be  cut  without 
imbedding.  The  two-  and  three-year  shoots  and  all  older  material 
should  be  cut  freehand,  without  imbedding,  and,  preferably,  before 
fixing.  Such  sections  are  transferred  directly  from  the  knife  to  95 
per  cent  alcohol. 

For  the  structure  of  the  adult  stem,  select  a  clear  board  and,  for 
transverse  sections,  cut  out  pieces  about  15  mm.  long  and  6  to  10  mm. 
square;  for  longitudinal  sections,  use  pieces  about  10  mm.  long,  with 
5  and  10  mm.  for  the  other  two  faces.  Cut  from  the  face  which  will 
give  sections  5X10  mm.  Orient  carefully,  so  that  the  longitudinal 
radial  sections  shall  be  exactly  parallel  with  the  rays,  and  the  longi- 
tudinal tangential  sections  exactly  tangential  to  the  rays.  Leave  the 
sections  in  95  per  cent  alcohol  for  15  or  20  minutes  before  staining. 
Stain  for  at  least  24  hours  in  safranin,  extract  the  stain  until  only  a 


Spermatophytes — Gymnosperms  253 

faint  red  color  is  left  in  the  cellulose  walls,  and  then  stain  in  Dela- 
field's  haematoxylin.  Stain  some  of  the  sections  in  safranin  and 
anilin  blue,  some  in  safranin  and  light  green,  and  some  in  iodine 
green  and  acid  fuchsin.  A  single  preparation  with  sections  stained 
in  various  ways  will  repay  a  careful  study.  Of  course,  every  prepara- 
tion should  contain  transverse,  longitudinal  radial  and  longitudinal 
tangential  sections. 

The  root. — The  primary  root  should  be  studied  in  the  embryo 
while  it  is  still  contained  in  the  seed.  Collect  material  in  September, 
October,  or  at  any  later  date.  If  material  is  collected  in  winter,  the 
seeds  should  be  soaked  in  water  for  a  day  or  two  before  fixing.  In 
any  case,  remove  the  testa  and  cut  a  thin  slab  from  opposite  sides 
of  the  endosperm  to  facilitate  fixing  and  infiltration.  For  secondary 
roots  and  also  for  the  structure  of  the  stele  in  the  primary  root, 
germinate  the  seeds  and  fix  material  after  the  hypocotyl  has  reached 
a  length  of  3  or  4  cm.  The  seeds  of  Pinus  edulis,  commonly  called 
Pinon,  or  edible  pine,  can  be  obtained  in  most  cities.  They  are 
particularly  good  for  a  study  of  the  mature  embryo  and  the  seedling. 

The  older  roots  are  treated  like  the  stems. 

The  leaves. — The  leaves  of  our  common  gymnosperms  cut  readily 
in  paraffin  while  they  are  young  and  tender,  but  as  they  approach 
maturity  'it  is  a  fruitless  task  to  attempt  paraffin  sections. 

Good  sections  may  be  obtained  in  great  quantities  with  little 
trouble  by  the  following  method:  Make  a  bunch  of  the  needles  as 
large  as  one's  little  finger,  wrap  them  firmly  together  with  a  string, 
allowing  about  |  inch  of  the  bunch  to  project  above  the  wrapping; 
then  fasten  the  whole  in  a  sliding  microtome  or  a  hand  microtome, 
and  every  stroke  of  the  razor  will  give  twenty  or  thirty  sections,  some 
of  which  will  surely  be  good.  As  the  sections  are  cut,  put  them  into 
95  per  cent  alcohol;  after  5  or  10  minutes,  transfer  to  70  per  cent 
alcohol,  where  they  should  remain  for  15  or  20  minutes  to  remove 
the  chlorophyll;  then  transfer  to  the  stain. 

Spermatogenesis.— In  October  the  clusters  of  staminate  cones 
which  are  to  shed  their  pollen  in  the  coming  spring  are  already  quite 
conspicuous.  The  cones  should  be  picked  off  separately,  and  the 
scales  should  be  carefully  removed  so  as  to  expose  the  delicate 


254  Methods  in  Plant  Histology 

greenish  cone  within.  At  this  time  the  sporogenous  cells  are  easily 
distinguished.  Material  collected  in  January,  or  at  any  time  before 
growth  is  resumed  in  the  spring,  shows  about  the  same  stage  of 
development.  If  it  is  desired  to  secure  a  series  of  stages  with  the 
least  possible  delay,  a  branch  bearing  numerous  clusters  of  cones 
may  be  brought  into  the  laboratory  and  placed  in  a  jar  of  water. 
Growth  is  more  satisfactory  in  case  of  branches  broken  off  in  the 
winter  than  in  those  brought  in  before  there  has  been  any  period  of 
rest.  The  material  can  be  examined  from  time  to  time,  and  a  com- 
plete series  is  easily  secured.  The  mitotic  figures  in  the  pollen 
mother  cells  furnish  exceptionally  instructive  preparations.  The 
two  mitoses  take  place  during  the  last  week  in  April  and  the  first 
week  in  May.  Staminate  cones  which  will  yield  mitotic  figures  can 
be  selected  with  considerable  certainty  by  examining  the  fresh 
material.  Crush  a  microsporangium  from  the  top  of  the  cone  and 
one  from  the  bottom,  add  a  small  drop  of  water  and  a  cover  to  each, 
and  examine.  If  there  are  pollen  tetrads  at  the  bottom,  but  only 
undivided  spore  mother  cells  at  the  top,  it  is  very  probable  that 
longitudinal  sections  of  the  cone  will  yield  the  figures.  If  a  drop  of 
methyl  green  be  allowed  to  run  under  the  cover,  it  will  enable  one 
to  see  whether  figures  are  present  or  not.  When  desirable  cones  are 
found  slabs  should  be  cut  from  two  sides,  in  order  that  the  fixing 
agent  may  penetrate  more  rapidly  and  that  infiltration  with  paraffin 
may  be  more  thorough. 

The  later  stages,  showing  the  germination  of  the  microspores, 
furnish  better  sections  if  the  cones  are  cut  transversely  into  small 
pieces  about  5  mm.  thick.  It  is  very  easy  to  get  excellent  mounts 
of  the  pollen  just  at  the  time  of  shedding,  which,  in  Pinus  Laricio 
in  the  vicinity  of  Chicago,  occurs  near  the  middle  of  June.  Shake 
a  large  number  of  cones  over  a  piece  of  paper,  thus  securing  an 
abundance  of  material;  then  transfer  to  formalin  alcohol.  With 
loose  pollen  grains,  there  is  a  great  loss  of  material  if  any  of  the 
chromo-acetic  acid  series,  with  the  attendant  washing,  is  used.  If 
the  material  is  so  abundant  that  plenty  will  be  left  after  all  the  loss, 
chromo-acetic  acid  may  be  used,  and  the  mitotic  figures,  which  may 
still  be  found,  are  likely  to  stain  more  brilliantly  than  after  formalin 


Spermatophytes — Gymnosperms  255 

alcohol.  However,  most  of  the  mitoses  take  place  before  the  pollen 
is  shed  or  after  it  reaches  the  nucellus.  Infiltration  in  the  bath  will 
not  require  more  than  30  minutes.  When  the  infiltration  is  com- 
plete, there  should  be  only  enough  paraffin  to  cover  the  mass  of  pollen 
grains.  The  material  may  now  be  poured  out  into  a  rectangular  dish 
or  box  with  surface  enough  to  make  the  cake  about  f  inch  thick. 
Good  results  may  be  secured  by  pouring  the  paraffin  upon  a  cold 
piece  of  glass.  Another  method  is  to  keep  the  material  in  a  small 
bottle  during  infiltration,  and  when  ready  to  imbed,  simply  cool  the 
bottle.  Break  the  bottle  carefully,  cut  off  the  lower  portion  of  the 
paraffin  containing  the  pollen,  mount  it  on  a  block  in  the  usual 
manner,  and  trim  away  some  of  the  paraffin  so  that  two  parallel 
surfaces  will  make  the  sections  ribbon  well.  Sections  should  not  be 
thicker  than  5  ju.  Material  in  this  stage  shows  a  large  tube  nucleus, 
a  somewhat  lenticular  (generative)  cell  with  a  more  deeply  staining 
nucleus,  and,  lastly,  two  small  prothallial  cells  quite  close  to  the  spore 
wall.  The  prothallial  cells  cannot  always  be  detected  at  this  stage, 
and  there  may  be  some  doubt  as  to  whether  two  such  cells  are  always 
present.  The  division  of  the  lenticular  cell  into  "stalk  cell"  and 
"body  cell,"  and  also  the  division  of  the  body  cell  into  the  two  male 
cells,  must  be  looked  for  in  sections  of  the  nucellus  of  the  ovule. 

Abies  balsamea  is  a  better  type  for  illustrating  spermatogenesis, 
since  the  pollen  mother  cells  and  the  pollen  grains  are  much  larger 
and  the  division  of  the  generative  cell  into  the  "stalk"  and  "body" 
cells  takes  place  before  the  pollen  is  shed  (Fig.  88). 

Araucaria  and  Agathis  are  the  best  forms  for  illustrating  numerous 
prothallial  cells.  Podocarpus  and  Taxodium  are  also  good.  Thuja  or 
Juniperus  may  be  used  to  illustrate  the  entire  absence  of  prothallial 
cells.  Both  Thuja  and  Juniperus  show  highly  organized  male  cells. 

Oogenesis.— In  Pinus  Larido  the  rudiment  of  the  ovulate  strobi- 
lus,  which  is  to  be  pollinated  in  June,  can  be  detected  in  the  preceding 
October.  The  collection  of  this  stage  is  very  uncertain,  because  there 
seems  to  be  no  mark  distinguishing  buds  containing  ovules  from  buds 
which  are  only  vegetative.  By  collecting  numerous  buds  from  the 
tops  of  vigorous  trees  which  are  known  to  produce  an  abundance  of 
strobili,  a  few  buds  containing  the  desired  stages  may  be  obtained. 


256 


Methods  in  Plant  Histology 


In  May,  after  the  strobili  break  through  the  bud  scales,  material 
is  easily  collected.  Up  to  the  time  of  pollination  the  entire  ovulate 
strobilus  cuts  easily  in  paraffin.  Longitudinal  sections  of  the  cone  at 
this  time  give  good  views  of  the  bract  and  ovuliferous  scale  bearing 
the  ovules.  The  integument  is  very  well  marked,  and  in  the  nucellus 
one  or  more  sporogenous  cells  can  usually  be  distinguished.  As  soon 
as  the  scales  close  up  after  pollination,  the  cone  begins  to  harden  and 
soon  makes  trouble  in  cutting.  Even  before  the  scales  close  up,  it 


Fia.  88. — Abies  balsamea:  photomicrograph  of  section  of  pollen  grain  about  the 
time  of  shedding;  there  are  two  prothallial  cells,  the  stalk  cell,  body  cell,  and  tube  cell; 
sometimes  the  division  in  the  body  cell  also  takes  place  before  the  pollen  is  shed;  from 
a  negative  by  Mr.  A.  H.  Hutchinson.  X815. 


is  better  to  cut  a  slab  from  opposite  sides  of  the  cone;  after  the  scales 
close,  it  is  almost  a  necessity.  For  sections  of  the  whole  cone,  fix 
in  formalin  alcohol.  Dr.  Hannah  Aase  succeeded  in  cutting  complete 
series  of  paraffin  sections  from  cones  of  Pinus  Banksiana  more  than 
2  cm.  in  length.  She  fixed  them  in  formalin  alcohol,  and  used  pro- 
longed periods  in  dehydrating,  clearing,  and  infiltrating.  Land's 
bichromate  of  potash  and  glue  fixative  was  used  in  fixing  the  sections 
to  the  slide.  Such  series  of  sections  of  large  cones  were  necessary  for 
an  investigation  of  the  vascular  anatomy. 


Spermatophytes — Gymnosperms 


257 


For  a  study  of  the  ovule  and  the  structures  within  it,  better 
preparations  will  be  obtained  by  carefully  cutting  off  the  pair  of 
ovules  from  the  scale :  Fix  in  chromo-acetic  acid  with  a  little  osmic 
acid.  In  free  nuclear  stages  of  the  female  gametophyte,  which  begin 
in  the  autumn,  are  interrupted  by  winter,  and  are  completed  in  May, 
plasmolysis  is  likely  to  occur.  After  walls  appear  there  is  less  danger. 
From  the  middle  of  May  to  the  first  of  July  collections  should  be 


A     B 


FIG    89—  Finns  Laricio:    A,  top  of  prothallium  with  an  archegonium  just  before 
the  cutting  off  of  the  ventral  canal  cell;  fixed  in  Flemming's  weaker  solution  and  stainc 
in  Haidenhain's  iron-alum  haematoxylin;    collected  June  18,  1897;    B,  C.  and  D 
stages  in  the  development  of  the  embryo;    fixed  in  chromo-acetic  acid  and  stained 
safranin,  gentian-violet,  orange;    collected  July  2,   1897.     > 

made  at  intervals  of  two  or  three  days,  since  during  these  six  weeks 
the  gametophyte  completes  the  free  nuclear  stage  and  develops 
cell  walls,  the  archegonium  completes  its  entire  development,  the 
egg  is  fertilized,  and  the  sporophyte  may  reach  the  suspensor  stage. 
For  preparations  like  that  represented  in  Fig.  89,  A,  it  is  a  good  plan 
to  remove  the  endosperm  with  its  archegonia  from  the  ovule, 
infiltration,  and  cutting  will  then  occasion  but  little  trouble  and  1 
whole  ribbon  may  be  got  upon  a  single  slide.     However,  at  this  stage 
the  pollen  tubes  with  their  contents  are  rapidly  working  their  way 


258 


Methods  in  Plant  Histology 


through  the  nucellus  toward  the  archegonia,  and  consequently,  in 
some  of  the  material,  it  is  better  to  retain  enough  of  the  tissues  of  the 
ovule  to  keep  the  nucellus  in  place.  In  later  stages,  after  fertilization 


PIG.  90. — Pinus  Laricio:  photomicrograph  showing  the  formation  of  the  ventral 
canal  cell;  usually,  this  cell  is  not  so  large  in  proportion  to  the  egg;  fixed  in  Flemming's 
weaker  solution  and  stained  in  safranin,  gentian-violet,  orange;  the  preparation  was  made 
in  1897,  the  photomicrograph  in  1915;  Cramer  contrast  plate;  4mm.  objective;  ocular 
X4;  Abb6  condenser;  yellowish-green  filter  and  also  a  strong  filter  used  in  outdoor 
photography;  camera  bellows,  75  cm.;  arc  light;  exposure,  7  minutes.  X587. 

has  taken  place,  the  developing  testa  should  be  removed  with  great 
care,  for  a  very  slight  pressure  is  sufficient  to  injure  the  delicate  parts 
within. 


Spermatophytes—Gymnosperms  259 

The  period  at  which  the  various  stages  may  be  found  varies 
with  the  species,  the  locality,  and  the  season.  In  Pinus  Laricio 
the  megaspore  mother  cells  appear  as  soon  as  the  young  strobili  break 

through  the  bud  scales.    At  Chicago,  in  the  season  of  1897,  material 

collected  May  27  did  not 

yet  show  archegonia;  the 

ventral  canal  cell  was  cut 

off  about  June  21   (see 

Fig.  90),  the  fusion  of  the 

egg    and    sperm    nuclei 

occurred   about  a  week 

later,    and     stages    like 

Fig.   89,  B,    C,   and   D, 

were  common  in  material 

collected  July  2.     In  the 

season  1896  all  the  stages 

appeared  about   two 

weeks  earlier.     In  Pinus 

sylvestris  the  stages  ap- 
peared   a    little    earlier 

than  in  Pinus  Laricio. 
After  the  stage  shown 
in  Fig.  89,  A,  has  ap- 
peared, it  is  necessary  to 
collect  every  day  until 
the  stage  shown  in 
Fig.  89,  D,  is  reached. 
If  collections  are  made  at 
intervals  of  3  or  4  days, 
the  most  interesting 
stages,  like  the  cutting 
off  of  the  ventral  canal 
cell,  fertilization,  and  the 
first  divisions  of  the  nucleus  of  the  oospore,  may  be  missed  altogether. 
It  should  be  mentioned  that  all  the  ovules  of  a  cone  will  be  in  very 
nearly  the  same  stage  of  development;  consequently  it  is  worth 


Fio.  91. — Pinus  Banksiana:  photomicrograph  of 
young  embryos  teased  out  by  the  method  described 
in  the  text;  from  a  preparation  by  Mr.  J.  T. 
Buchholz;  Cramer  contrast  plate;  16  mm.  objective; 
no  ocular  or  Abb£  condenser;  camera  bellows.  75cm.; 
safranin  filter;  arclight;  exposure,  17 seconds.  X54. 


260  Methods  in  Plant  Histology 

while  to  keep  the  ovules  from  each  separate.  Stages  like  that  shown 
in  Fig.  90  are  rare  in  miscellaneous  collections,  but  if  ovules  from 
each  cone  are  kept  separate  and  this  figure  is  found,  the  rest  of 
the  ovules  from  that  cone  will  be  likely  to  show  some  phase  of  this 
interesting  mitosis. 

Thuja  and  Juniperus  are  good  types  to  illustrate  the  arche- 
gonium  complex  and  the  large,  highly  organized  male  cells.  In 
Thuja  a  series  from  the  appearance  of  archegonium  initials  to 
young  embryos  may  be  collected  between  June  10  and  June  20.  In 
Juniperus  pollination  occurs  late  in  May  and  fertilization  takes 
place  12|  months  later.  The  megaspores  are  formed  late  in  April 
and  the  development  of  the  female  gametophyte  occupies  about  6 
weeks. 

The  Embryo. — The  early  stages  of  the  sporophyte,  usually 
designated  as  the  proembryo,  have  been  mentioned  already. 

From  the  time  when  the  suspensors  begin  to  elongate  up  to  the 
appearance  of  cotyledons,  instructive  preparations  can  be  made  by 
mounting  the  embryo  whole.  Mr.  J.  T.  Buchholz  has  developed  a 
method  for  handling  these  small  objects.  Remove  the  testa  and 
then,  under  water,  hold  the  endosperm  gently  with  forceps  and 
press  the  neck  and  upper  part  of  the  archegonium  with  a  needle, 
pressing,  and  at  the  same  time  drawing  the  needle  away,  so  as  to  pull 
the  young  embryo  out.  Some  of  the  embryos  will  be  broken,  but 
by  careful  manipulation  more  than  half  should  be  entirely  uninjured. 
Fix  in  formalin  (5  per  cent  in  water) ,  stain  in  Delafield's  haematoxy- 
lin,  transfer  to  10  per  cent  glycerin,  and  continue  with  the  Venetian 
turpentine  method.  A  preparation  made  in  this  way  is  shown  in 
the  photomicrograph,  Fig.  91. 

These  stages,  and  all  subsequent  stages,  are  easily  cut  in  paraffin 
without  removing  the  embryo  from  the  endosperm.  Cut  a  thin 
slab  from  opposite  sides  of  the  endosperm,  fix  in  chromo-acetic  acid, 
with  or  without  a  little  osmic  acid,  imbed  in  paraffin,  and  stain  in 
safranin  and  gentian-violet.  This  will  give  a  good  view  of  the  abun- 
dant starch  and  other  food  stuff,  and  at  the  same  time  will  bring  out 
sharply  the  cell  walls  of  the  embryo. 


CHAPTER  XXV 

SPERMATOPHYTES 

ANGIOSPERMS 

This  group  is  so  large,  and  its  structures  are  so  varied  and  complex 
that  great  care  must  be  taken  in  the  selection  of  material  for  sections.' 
With  experience,  one  will  gradually  learn  what  stages  in  floral 
development,  what  stages  in  the  development  of  the  embryo-sac, 
or  what  stages  in  spermatogenesis  are  likely  to  be  correlated  with 
easily  recognized  field  characters. 

The  Vegetative  Structures.— In  stems,  roots,  and  leaves  the 
more  delicate  structures  should  be  imbedded  in  paraffin  and  the  more 
rigid  structures  should  be  cut  without  imbedding  at  all. 

The  stem. — Throughout  the  angiosperms,  the  vascular  cylinder 
is  an  endarch  siphonostele.  For  a  study  of  the  development  of  the 
stem,  the  common  geranium  (Pelargonium)  may  be  recommended. 
Near  the  base  of  a  fresh  stem,  about  1  cm.  in  diameter,  cut  freehand 
sections  and  fix  them  in  35  per  cent  alcohol  for  10  to  20  minutes; 
transfer  to  70  per  cent  alcohol  to  extract  the  chlorophyll,  and  then 
stain  in  safranin  and  light  green.  Such  sections  will  show  both 
primary  and  secondary  structures  in  the  stele  and  in  the  cortex. 
Higher  up,  there  will  be  secondary  structures  only  in  the  stele;  and 
still  higher  up  will  be  found  the  origin  of  interfascicular  cambium. 
All  these  can  be  cut  without  imbedding,  but  the  earlier  stages  show- 
ing the  differentiation  of  protoxylem,  metaxylem,  and  the  origin  of 
secondary  xylem  are  too  soft  for  successful  freehand  sections.  Fix 
in  formalin  alcohol  and  imbed  in  paraffin. 

For  a  study  of  woody  stems,  Tilia  americana  (basswood)  is  good, 
and  shoots  from  5  to  10  mm.  in  diameter  are  easy  to  cut.  Very  hard 
stems  like  Hicoria  (hickory)  and  Quercus  (oak)  must  be  boiled  and 
treated  with  hydro-fluoric  acid,  if  you  expect  to  cut  shoots  more  than 
5  to  7  mm.  in  diameter.  However,  with  a  good  sharp  knife  and  a 
rigid  microtome  much  larger  sections  can  be  cut  without  resorting 

261 


262 


Methods  in  Plant  Histology 


to  hydrofluoric  acid.     Of  course,  veneer  machines  cut  very  large  and 
fairly  thin,  smooth  sections  from  the  most  refractory  woods. 

While  a  random  selection  of  stems  would  furnish  material  for 
practice  in  technic,  we  suggest  that  the  stem  of  Clintonia  shows  a  good 
siphonostele  in  a  monocotyl;  the  rhizome  of  Acorus  calamus  is  a 
good  type  for  the  amphivasal  bundle;  Zea  Mays  shows  scattered 

bundles,  but  not  the  amphi- 
vasal condition;  Aloe  illus- 
trates secondary  wood  in 
monocotyls;  Iris  has  a  highly 
developed  endodermis  in  the 
rhizome;  and  Nymphea  or 
Nuphar  will  show  scattered 
bundles  in  a  dicotyl. 

The  sieve  tubes  of  the  phloem 
are  easily  demonstrated 
in  Cucurbita  Pepo,  the  com- 
mon pumpkin;  other  mem- 
bers of  the '  family  furnish 
good  material.  Take  pieces 
of  stem  about  1  cm.  long  and 
not  too  hard  to  cut  in  paraffin, 
fix  in  formalin  alcohol,  and 
stain  in  safranin,  gentian- 
violet,  orange.  The  tropical 
Tetracera,  one  of  the 


iron-alum   haematoxylin.     Cramer  contrast          that  they  are  easily  Seen  with 
plate;  4  mm.  objective;  ocular  X4;  camera  .  .  _,  , 

bellows.  85  cm.;    yellowish-green  filter  and  a    pocket    lens.       The    phloem 

™krrPn(iuTuuiu«£.us^outdoor  *™  «• «"  ^° fc th« iare<* 

stems  that  it  can  be  cut  out 

for  imbedding  in  paraffin  long  after  the  entire  stem  has  become 
too  hard  for  paraffin  sections. 

Roots. — It  has  long  been  known  that  the  root-tip  furnishes 
constantly  available  material  for  a  study  of  mitosis  (Fig.  92).  An 
onion  thrown  into  a  pan  of  water  will  soon  send  out  numerous  roots. 


Spermatophytes—Angiosperms  263 

Soak  beans  in  water  for  several  hours,  and  then  plant  them  about 
an  inch  deep  in  loose,  moist  sawdust.  The  primary  root  will  be 
long  enough  in  2  or  3  days.  The  large,  flat  beans  are  best.  Vicia 
Faba  is  very  favorable.  The  root-tips  of  Tradescantia  virginica, 
Ins  versicolor,  Podyphyllum  peltatum,  Arisaema  tryphyllum,  Cypri- 
pedium  pubescens,  and  many  others  furnish  excellent  material. 

Doubtless  cell  division  does  not  proceed  with  equal  rapidity 
at  all  hours  of  the  day.  Kellicott1  has  shown  that  in  the  root- 
tips  of  Allium  there  are  in  each  24  hours  two  periods  at  which 
cell  division  is  at  the  maximum,  and  two  at  which  it  is  at 
the  minimum.  The  maximum  periods  are  shortly  before  midnight 
(11 :00  P.M.),  and  shortly  after  noon  (1 :00  P.M.).  The  minima,  when 
cell  division  is  at  the  lowest  ebb,  occur  about  7:00  A.M.  and  3:00  P.M. 
When  cell  division  is  most  vigorous,  there  is  little  elongation,  and 
when  cell  division  is  at  the  minimum,  cell  elongation  is  at  the  maxi- 
mum. Consequently,  root-tips  of  Allium  should  be  collected  about 
1:00  P.M.  or  11: 00  P.M. 

We  have  not  made  any  systematic  series  of  experiments  to  test 
Kellicott's  results,  but  miscellaneous  observations  seem  to  indicate 
that  his  claim  holds  good  for  root-tips.  It  is  certain,  however,  that 
abundant  mitoses  may  be  found  at  other  times — even  at  3:00  P.M. — 
in  sporangia  of  ferns,  in  anthers  of  angiosperms,  in  endosperm,  and 
in  free  nucleus  stages  of  the  embryo  of  gymnosperms. 

Mitotic  figures  play  such  an  important  part  in  the  development 
of  the  plant  and  in  modern  theories  of  heredity,  that  it  is  worth  while 
to  acquire  a  critical  technic  in  fixing  and  staining  these  structures. 
Use  the  various  fixing  agents— Flemming's  weaker  solution,  chromo- 
acetic  acid  with  or  without  a  little  osmic,  Benda's  fluid,  Bouin's 
fluid,  corrosive  sublimate  with  acetic  acid,  and  any  others.  Make 
yourself  master  of  Haidenhain's  iron-alum  haematoxylin;  then  add 
the  safranin,  gentian- violet,  orange  combination;  then  safranin  and 
anilin  blue;  and  then  experiment  for  yourself,  but  remember  that  the 
triumphs  of  modern  cytology  have  been  won  with  iron-haematoxylin 
and  that  you  cannot  read  intelligently  the  literature  of  the  past  two 

»  Kellicott,  W.  E.,  "The  Daily  Periodicity  of  Cell  Division  and  of  Elongation  in  the 
Root  Of  Allium,"  Bull.  Torrey  Bot.  Club  31:529-550,  1904. 


264  Methods  in  Plant  Histology 

decades  until  you  have  gained  at  least  an  approximate  mastery 
of  this  stain.  Of  course,  dehydration,  clearing,  and  infiltration  must 
be  very  gradual.  The  schedules  by  Yamanouchi  and  by  Sharp,  on 
pp.  42  and  43,  will  repay  careful  study. 

In  staining  with  safranin,  gentian-violet,  orange,  allow  the 
alcoholic  safranin  to  act  for  16  to  24  hours;  then  extract  it  with  50 
per  cent  alcohol,  slightly  acidulated  with  hydrochloric  acid,  if  neces- 
sary, until  the  stain  has  almost  disappeared  from  the  spindle;  then 
pass  through  70,  85,  95,  and  100  per  cent  alcohol;  stain  in  gentian- 
violet  dissolved  in  clove  oil,  or  in  a  mixture  of  clove  oil  and  absolute 
alcohol,  for  5  to  20  minutes;  follow  with  orange  dissolved  in  clove 
oil,  remembering  that  this  will  weaken  the  safranin  and  sometimes 
the  gentian-violet;  finally  use  pure  clove  oil  to  differentiate  the 
gentian-violet.  Leave  the  slide  in  xylol  for  2  to  5  minutes  to  remove 
the  clove  oil  and  to  hasten  the  hardening  of  the  balsam. 

The  structure  and  development  of  the  young  root  will  be  shown, 
to  some  extent,  in  preparations  made  for  mitotic  figures.  The  origin 
of  dermatogen,  periblem,  plerome,  and  also  of  protoxylem,  is  well 
shown  in  Zea  Mays.  An  ear  of  sweet  corn,  as  young  and  tender  as 
you  can  find  on  the  market,  will  furnish  material.  Cut  out  from  the 
grain  a  rectangular  piece  about  2X3  mm.  and  4  or  5mm.  long;  if 
you  want  to  show  also  the  structure  of  the  entire  grain,  take  a  section 
the  entire  length  of  the  grain,  perpendicular  to  the  flat  side  of  the 
grain,  and  about  2mm.  wide.  Cut  the  latter  longitudinally;  the 
rectangular  pieces  are  sufficient  for  transverse  sections.  Fix  in 
chromo-acetic  acid. 

The  roots  of  Ranunculus  repens  furnish  good  illustrations  of  the 
radial  arrangement  of  xylem  and  phloem.  Smilax  shows  the  radial 
arrangement,  with  a  large  number  of  poles  and  a  very  highly  differ- 
entiated endodermis.  The  origin  of  secondary  xylem  and  phloem  is 
well  shown  in  Sambucus  nigra.  Vicia  Faba  shows  very  clearly  the 
origin  of  secondary  roots.  The  arrangement  of  cells  in  the  young 
roots  of  aquatic  or  semi-aquatic  plants  is  often  extremely  regular 
(Fig.  93). 

The  Leaf. — Young  and  tender  leaves  should  be  fixed  in  formalin 
alcohol  and  cut  in  paraffin.  Cut  sections  freehand  whenever  there 


Spermatophytes—Angiosperms  265 

is  sufficient  rigidity.  Resort  to  pith  only  when  necessary  In 
cutting  sections  of  a  leaf  like  that  of  Lilium,  lay  one  leaf  on  another 
until  you  have  a  bundle  of  them  which  will  be  nearly  square  in  trans 
verse  section.  Wrap  the  bundle  with  string  for  about  15  mm  -  cut 
the  bundle  transversely  so  that  about  5  mm.  of  the  bundle  will 
project  beyond  the  tied  portion.  Fasten  the  tied  portion  in  the 


FIG.  93. — Sparganium  eurycarpum:  photomicrograph  of  transverse  section  of  young 
root:  fixed  in  chromo-acetic  acid  and  stained  in  Bismarck  brown;  Cramer  contrast  plate; 
16  mm.  objective;  ocular  X4;  no  Abbfi  condenser;  yellowish-green  filter;  camera  bel- 
lows, 1  meter;  exposure,  8  seconds.  X90. 

microtome,  or  hold  it  in  your  fingers,  and  cut  transverse  sections. 
About  15  to  20/i  is  a  good  thickness  for  general  leaf  structure. 
The  sections  of  leaves  in  the  center  of  such  a  bundle  are  likely 
to  be  better  than  you  could  cut  from  single  leaves  held  in  pith. 
In  case  of  large  leaves,  cut  out  strips  about  1  cm.  square  and  tie 
them  together. 

Buds  will  furnish  beautiful  preparations  of  young  leaves  and,  at 
the  same  time,  will  show  the  vernation.  Cut  the  bud  transversely, 


266  Methods  in  Plant  Histology 

a  little  above  the  middle;  remove  the  bud  scales,  if  they  promise 
to  cause  trouble;  retain  only  enough  tissue  at  the  base  of  the  bud 
to  hold  the  parts  in  place.  Fix  in  formalin  alcohol  and  stain  in 
safranin  and  light  green. 

Epidermis  stripped  from  the  leaf,  fixed  in  10  per  cent  formalin 
in  water  for  a  day  or  two,  and  then  stained  in  safranin  and  anilin 
blue,  will  give  excellent  views  of  stomata.  The  development  of 
stomata  is  particularly  well  shown  in  Sedum  purpurascens,  even  in 
leaves  which  have  reached  the  adult  size.  The  epidermis  is  very 
easily  stripped  from  a  leaf  of  Sedum. 

Floral  Development. — For  a  study  of  floral  development  very 
young  buds  are  necessary,  and  it  is  best  to  select  those  forms  which 
have  rather  dense  clusters  of  flowers,  in  order  that  a  complete  series 
may  be  obtained  with  as  little  trouble  as  possible. 

The  usual  order  of  appearance  of  floral  parts  is  (1)  calyx,  (2) 
corolla,  (3)  stamens,  and  (4)  carpels;  but  if  any  of  these  organs  is 
reduced  or  metamorphosed,  their  order  of  appearance  may  be 
affected. 

Floral  development  is  easily  studied  in  the  common  Capsella 
bursa-pastoris.  The  best  time  to  collect  material  is  late  in  March 
or  early  in  April.  Dig  up  the  plant,  carefully  remove  the  leaves, 
and  in  the  center  of  the  rosette  a  tiny  white  axis  will  be  found.  A 
series  of  these  axes  from  3  to  9  mm.  in  length,  and  from  1 . 5  to 
3 . 5  mm.  in  diameter  will  give  a  very  complete  series  of  stages  in 
the  development  of  the  floral  organs.  Preparations  from  the  apex 
of  the  shoot  taken  after  the  inflorescence  appears  above  ground  are 
not  to  be  compared  with  those  taken  early  in  the  season,  because 
the  pedicels  begin  to  diverge  so  early  that  median  longitudinal  sec- 
tions of  the  flowers  are  comparatively  rare.  Fix  in  chromo-acetic 
acid  and  stain  in  Delafield's  haematoxylin.  The  sections  should  be 
longitudinal  and  about  5  /i  thick.  Capsella  shows  the  hypogynous 
type  of  development.  The  order  of  appearance  of  floral  parts  is 
(1)  calyx,  (2)  stamens,  (3)  carpels,  and  (4)  petals.  The  ovary  is 
compound  (syncarpous) . 

Ranunculus,  which  is  also  hypogynous,  will  illustrate  the  develop- 
ment of  the  simple  (apocarpous)  ovary.  The  ovules  appear  quite 


Spermatophytes — Angiosperms  267 

early,  so  that  the  archesporial  cell,  or  even  the  megaspores,  may  be 
seen  while  the  carpel  is  still  as  open  as  in  any  gymnosperm. 

In  the  willows,  Salix,  the  bud  scales  must  be  removed  and  the 
copious  hairs  should  be  trimmed  off  as  much  as  possible  with  scissors, 
after  which  the  catkin  should  be  cut  in  two  longitudinally  and  placed 
in  the  fixing  agent. 

The  cat-tail,  Typha,  presents  a  simple  type  of  floral  development. 
The  leaves  should  be  dissected  away  long  before  the  flowers  can  be 
seen  from  the  outside.  The  cylindrical  clusters,  varying  in  diameter 
from  2  or  3  mm.  up  to  the  size  of  one's  finger,  will  afford  a  complete 
series  of  stages.  Until  the  spadix  reaches  the  diameter  of  a  lead 
pencil,  transverse  sections  are  easily  cut.  For  later  stages,  the  outer 
part  of  the  spadix  should  be  sliced  off  so  that  only  enough  spadix  is 
retained  to  hold  the  florets  in  place. 

Prunus  and  many  other  members  of  the  Rosaceae  furnish 
examples  of  the  perigynous  type  of  development.  In  many  of  them 
the  floral  parts  do  not  occur  in  the  usual  succession. 

The  epigynous  type  is  well  shown  in  the  Compositae.  The  order 
of  appearance  is  (1)  corolla,  (2)  stamens,  (3)  carpels,  and  (4)  calyx 
(pappus). 

The  common  dandelion,  Taraxacum  offianale,  affords  an  excellent 
series  with  little  labor.  Examine  vigorous  plants  which  have,  as  yet, 
no  flowers  or  buds  in  sight.  Dig  up  the  plant  and  dissect  away  the 
leaves.  If  there  is  a  white  cluster  of  flower  buds,  the  largest  not 
more  than  4  mm.  in  diameter,  cut  out  the  cluster,  leaving  only 
enough  tissue  at  the  base  to  hold  the  buds  in  place.  Larger  heads 
should  be  cut  separately. 

Our  most  common  thistle,  Cirsium  lanceolatum,  shows  the  floral 
development  with  unusual  clearness,  but  the  preparation  of  the 
material  is  somewhat  tedious.     The  involucre,  which  is  too  hard  to 
cut,  must  be  carefully  dissected  away.     Retain  only  enough  of  the 
receptacle  to  hold  the  developing  florets  in  place.     A  series  of  s 
with  disks  varying  from  3  mm.  to  10  mm.  in  diameter  will  show  tl 
development  from  the  undifferentiated  papilla  up  to  the  appcaranc 
of  the  archesporial  cell  in  the  nucellus  of  the  ovule.     The  Canada 
thistle,  Cirrium  arvense,  is  equally  good,  but  it  is  more  difficult 


268  Methods  in  Plant  Histology 

dissect  out  the  desirable  parts.  In  the  common  sunflower,  Helian- 
thus  annuus,  the  young  floral  parts,  like  the  mature  head,  are  so  very 
large  that  a  satisfactory  study  may  be  made  with  a  low-power 
objective.  As  in  case  of  the  thistle,  the  involucre  must  be  trimmed 
away  and  only  enough  of  the  receptacle  retained  to  hold  the  florets 
together. 

Spermatogenesis. — The  earlier  stages  in  spermatogenesis  will 
be  found  in  the  preparations  of  floral  development.  The  origin  of 
the  archesporium,  the  origin  of  sporogenous  tissue,  and  the  formation 
of  the  tapetum  are  beautifully  shown  in  longitudinal  and  in  transverse 
sections  of  the  anthers  of  Taraxacum  and  many  other  Compositae. 
Transverse  sections  of  the  head  of  Taraxacum  or  any  similiar  head 
at  the  time  when  pollen  mother  cells  are  rounding  off  in  the  center  of 
the  head,  will  show  various  stages  from  the  mother  cells  in  the  center 
to  the  tetrads  of  spores  at  the  periphery.  Transverse  sections  of 
the  anther  of  Polygala  give  exceptionally  well-defined  views  of  the 
archesporial  cells  and  sporogenous  areas. 

Lilium,  Trillium,  Galtonia,  Iris,  Tradescantia,  Vicia,  and  Podo- 
phyllum  can  be  recommended  for  demonstrating  the  nuclear  changes 
involved  in  the  formation  of  spores  from  the  mother  cell  (Fig.  94). 
Several  species  of  Lilium  are  common  in  greenhouses,  and  these  may 
be  used  where  wild  material  is  not  available.  In  early  stages,  where 
the  sporogenous  cells  have  not  yet  begun  to  round  off  into  spore 
mother  cells,  it  is  sufficient  to  remove  the  perianth,  retaining  just 
enough  of  the  receptacle  to  hold  the  stamens  in  place.  Transverse 
sections  show  the  six  stamens  and  also  the  young  ovary.  After  the 
spore  mother  cells  have  begun  to  round  off,  each  stamen  should  be 
removed  so  as  to  be  cut  separately.  In  securing  the  desirable  stages 
showing  the  division  of  the  mother  cell  into  microspores,  much  time 
and  patience  will  be  saved  by  determining  the  stage  of  development 
before  fixing  the  material.  Mitosis  is  more  or  less  simultaneous 
throughout  an  anther.  Long  anthers  are  particularly  favorable, 
since  they  may  show  a  very  closely  graded  series  of  the  various  phases 
of  mitosis.  An  anther  of  Iris  may  show  mother  cells  with  nuclei  in 
synapsis  at  the  top,  while  the  mother  cells  at  the  bottom  have  reached 
the  equatorial  plate  stage  of  the  first  division;  or,  the  mother  cells 


Spermatophytes—Angiosperms  269 

at  the  top  may  show  the  first  division,  while  those  at  the  bottom 
show  the  second.  Determine  the  stage  by  examining  a  few  mother 
cells  before  fixing. 

From  what  has  been  said,  it  is  evident  that  longitudinal  sections 
should  be  cut  to  show  mitosis.  Transverse  sections  should  be  cut 
to  show  the  general  structure  of  the  anther.  It  is  not  necessary  to 


FIG.  94. — Lilium  candidum:  photomicrograph  of  mitosis  in  pollen  mother  cells; 
in  one  of  the  pollen  mother  cells  the  twelve  chromosomes  can  be  counted ;  from  a  prepara- 
tion by  F.  L.  Pickett.  X260. 

cut  the  stamens  into  pieces  before  fixing,  since  they  are  easily  pene- 
trated and  infiltrated;  in  later  stages  the  stamens  must  not  be  cut 
into  pieces,  since  the  pollen  grains  are  easily  washed  out. 

The  problem  of  fixing  spore  mother  cells  has  received  much  atten- 
tion. In  fixing  mother  cells  and  the  two  mitoses  by  which  spores 
are  formed,  investigators  have  used  almost  exclusively  the  chromo- 
osmo-acetic  acid  solutions  of  Flemming,  some  preferring  the 
weaker  solution  and  some  the  stronger.  These  solutions  have 
been  used  in  nearly  all  of  the  work  of  the  Bonn  (Germany)  school. 


270  Methods  in  Plant  Histology 

Osterhout1  experimented  with  forty  fixing  agents,  and  then  con- 
cluded that  Flemming's  stronger  solution  was  the  best.  Professor 
Gregoire  and  his  students  have  made  this  their  principal  fixing  agent. 
In  spite  of  the  weight  of  authority,  we  believe  that  the  value  of 
solutions  with  such  a  large  proportion  of  osmic  acid  has  been  over- 
estimated. Some  osmic  acid  is,  doubtless,  desirable,  but  we  should  use 
only  half  the  amount  of  osmic  recommended  in  Flemming's  weaker 
solution.  The  formula  for  that  solution  is  often  given  as  follows: 

(  Chromic  acid  (1  per  cent) -25  c.c. 

A  \  Glacial  acetic  acid  (1  per  cent) 10  c.c. 

I  Water 55  c.c. 

B.  Osmic  acid  (1  per  cent) 10  c.c. 

Keep  the  mixture  A  made  up,  and  add  B  as  the  reagent  is  needed 
for  use,  since  the  solution  does  not  keep  well.  One  seldom  uses  this 
reagent  in  large  quantities.  About  40  c.c.  is  as  much  as  one  is  likely 
to  need  for  any  collection  of  anthers  or  root-tips.  Take  36  c.c.  of  A 
and  4  c.c.  of  B.  It  will  be  worth  while  to  try  36  c.c.  of  A  and  2  c.c. 
of  B,  or  even  1  c.c.  of  B.  If  the  regular  formula  is  used,  we  should 
let  it  act  for  an  hour,  and  then  replace  it  by  A,  without  any  osmic 
acid.  The  osmic  acid  undoubtedly  accelerates  the  killing  of  the 
protoplasm.  This  is  seen  more  readily  in  animals.  If  Cyclops  be 
brought  into  30  c.c.  of  the  solution  A,  the  animals  will  swim  for 
awhile;  if  5  or  6  drops  of  1  per  cent  osmic  acid  be  added  to  the 
solution,  the  animals  cease  their  movements  almost  instantly. 
Doubtless  the  osmic  acid  has  the  same  effect  upon  plant  protoplasm. 
Where  fixing  is  slow,  very  few  mitotic  figures  are  found  with  the 
chromosomes  midway  between  the  equator  and  the  poles.  The 
addition  of  10  drops  of  1  per  cent  osmic  acid  to  50  c.c.  of  the  solution 
just  mentioned  will  secure  as  large  a  proportion  of  anaphases  as 
solutions  which  are  stronger  in  osmic  acid,  and  there  is  no  disagreeable 
blackening. 

Farmer  and  Shove,2  in  studying  these  mitoses  and  also  vegeta- 
tive mitoses  in  Tradescantia,  secured  better  results  with  a  mixture 

1  Osterhout,  W.  J.  V.,  "Cell  Studies,  I,  Spindle  Formation  in  Agave,"  Proc.  Cal. 
Acad.  Sci.  Botany,  Third  Series,  2:255-284,  1902. 

2  Farmer,  J.  B.,  and  Shove,  Dorothy,  "On  the  Structure  and  Development  of  the 
Somatic  and  Heterotype  Chromosomes  of  Tradescantia  virginica,"  Quart.  Jour.  Mic.  Sci. 
48:559-569,  1905. 


Spermatophytes—Angiosperms  271 

of  2  parts  of  absolute  alcohol  and  1  part  glacial  acetic  acid  They 
allowed  the  fixing  agent  to  act  15  to  20  minutes,  then  washed  in 
absolute  alcohol,  and  imbedded  by  the  usual  methods.  This  propor- 
tion of  acetic  acid  seems  entirely  too  large  for  any  accurate  work 
with  chromatin,  and  we  doubt 
whether  the  structure  of  the 
cytoplasm  is  normal  when  so 
much  acetic  acid  is  used. 

The  entire  pollen  mother 
cell  may  be  stained  and  mounted 
without  sectioning.  Two  de- 
scriptions of  technic  appeared 
in  1912,  one  by  Mann1  and  the 
other  by  Pickett.2  Mann  re- 
moves the  pollen  mother  cells 
before  fixing  and  staining; 
Pickett  fixes  and  stains  the 
anther  in  toto  and  teases  out 
the  pollen  mother  cells  just 
before  mounting. 

In  Mann's  method,  the 
anther  is  placed  in  a  drop  of 
water  and  the  tip  is  cut  off;  a 
gentle  tapping  with  a  needle 
will  then  cause  the  pollen  mother 
cells  to  float  out  into  the  drop. 
Fix  in  Bouin's  fluid,  4  to  8  hours, 
wash  in  50  per  cent  alcohol 
until  no  color  remains,  and  then 
stain  in  iron-haematoxylin.  At 
this  stage  we  should  put  the  material  into  10  per  cent  glycerin  and 
follow  the  Venetian  turpentine  method. 

Pickett  fixed  entire  anthers  in  chromo-acetic  acid  for  30  hours, 
washed  in  water  for  24  hours,  and  then  passed  up  to  80  per  cent 

'Mann,  Albert,  "The  Preparation  of  Unbroken  Pollen  Mother  Cells  and  Other 
Cells  for  Studies  in  Mitosis,"  Science,  36:  151-153.  1912. 

2  Pickett,  P.  L.,  "Preparation  of  Whole  Pollen  Mother  Cells,"  Sriencr,  36:  479-480. 
1912. 


FIG.  95.  —  Erythronium  americanum: 
photomicrograph  of  mature  pollen  grains; 
the  one  at  the  top,  which  is  cut  longitudin- 
ally, shows  both  the  tube  nucleus  and  the 
conspicuous  generative  cell;  the  other  is 
cut  transversely  and  shows  the  generative 
cell,  but  not  the  tube  nucleus;  stained  in 
safranin  and  gentian-violet;  from  a  prep- 
aration by  Dr.  Lula  Pace;  Cramer  con- 
trast plate;  4  mm.  objective;  ocular  X4; 
yellowish-green  filter;  bellows,  85  cm.; 
exposure,  3  minutes.  X015. 


272 


Methods  in  Plant  Histology 


alcohol.  At  this  point,  he  stained  in  strong  cochineal  or  Kleinenberg's 
haematoxylin  for  5  days,  then  completed  the  dehydration,  cleared 
in  cedar  oil,  teased  out  the  mother  cells,  and  mounted  in  balsam. 
In  dealing  with  the  whole  anther,  it  is  necessary  to  select  stains 
which  will  not  overstain.  Alum  cochineal  and  Mayer's  haem-alum 
might  be  suggested.  It  would  be  worth  while  to  try  a  combination 


FIG.  96. — Lilium  philadelphicum:  photomicrograph  of  section  of  young  ovule  showing 
the  conspicuous  archesporial  cell;  fixed  in  chromo-acetic  acid  and  stained  in  Delafleld's 
haematoxylin  and  erythrosin.  X308. 

of  the  two  methods.  Fix  the  entire  anther  in  chromo-acetic  acid, 
wash  in  water,  and  then  stain  in  iron-haematoxylin.  When  the  last 
stage  in  staining  is  reached — the  extraction  of  the  stain  in  iron- 
alum — remove  the  pollen  mother  cells  and  watch  the  differentiation; 
then  wash  in  water  and  follow  the  Venetian  turpentine  method. 

The  pollen  grain  at  the  time  of  shedding  generally  consists  of  two 
cells,  the  tube  cell  and  the  generative  cell,  which  afterward  divides 


Spermatophytes—A  ngiosperms 


273 

and  forms  two  male  cells  or  two  male  nuclei.  Lilium  and  Ery- 
thronium  furnish  good  illustrations  of  pollen  shed  in  the  two-cell 
stage  (Fig.  95).  In  Silphium,  Sambucus,  and  Sagittaria  the  genera- 
tive  nucleus  divides  before  the  pollen  is  shed. 

Sections  should  not  be  more  than  3  to  5  /*  thick,  if  they  are  to 
show  a  clear  differentiation  of  exine,  intine,  starch,  and  other  struc- 
tures. If  sections  have  been  stained  in  iron-haematoxylin,  staining 


FIG.  97. — Lilium  philadelphicum:  photomicrograph  of  transverse  section  of  ovary 
showing,  in  one  of  the  ovules  on  the  left,  the  first  mitosis  in  the  megaspore  mother  cell ; 
and,  in  one  of  the  ovules  on  the  right,  the  second  mitosis  which  gives  rise  to  the  four 
megaspore  nuclei — chromo-acetic  acid;  safranin,  gentian-violet,  orange.  Cramer  con- 
trast plate;  16  mm.  objective;  ocular  X4;  yellowish-green  filter  and  also  a  strong  filter 
such  as  is  used  in  outdoor  work;  camera  bellows,  30  cm. ;  exposure,  2  minutes.  X64. 


in  safranin  for  from  3  to  7  minutes  will  give  the  exine  a  bright-red 
color  and  will  not  obscure  the  haematoxylin.  A  rather  sharp  stain 
in  gentian-violet  will  stain  the  starch  and  also  the  intine.  In 
Asclepias  and  many  orchids,  in  which  a  common  exine  surrounds  the 
entire  mass  of  pollen  grains,  care  must  be  taken  not  to  overstain. 

In  many  cases  the  pollen  grains  will  put  out  their  tutas  in  a  2  to 
5  per  cent  solution  of  cane-sugar  in  water.     Where  the  interval 


274 


Methods  in  Plant  Histology 


between  pollination  and  fertilization  is  known  (about  72  hours  in 
Lilium),  pieces  of  the  stigma  and  style  showing  pollen  tubes  can  be 
selected  with  some  certainty. 

Ob'genesis. — As  in  spermatogenesis,  the  early  stages  will  be  found 
in  preparations  of  floral  development.     The  preparations  of  Capsella 


Fia.  98. — Lilium  philadelphicum:  photomicrograph  of  second  mitosis  in  megaspore 
mother  cell — chromo-acetic  acid;  safranin,  gentian-violet,  orange.  Cramer  contrast 
plate;  4mm.  objective;  ocular  X4;  Abbe  condenser  camera  bellows,  1  meter;  yellowish- 
green  filter  and  also  a  strong  filter  such  as  is  used  in  outdoor  work;  camera  bellows, 
1  meter;  exposure,  7  minutes.  X626. 

will  show  the  origin  and  development  of  the  nucellus  (megaspo- 
rangium)  and  also  the  megaspore  mother  cell.  The  division  of  the 
megaspore  mother  cell  to  form  four  megaspores  takes  place  shortly 
before  the  bud  begins  to  unfold.  A  massive  megasporangium  with 


Spermatophytes — Angiosperms  275 

several  megaspore  mother  cells  may  be  found  in  Ranunculus;  a 
megasporangium  with  only  one  megaspore  mother  cell  and  only  one 
layer  of  cells  surrounding  it  may  be  found  in  any  of  the  Compositae. 
Senecio  aureus  and  Erectites  hieracifolium  are  good  and  are  particu- 
larly easy  to  cut.  In  Trillium  and  in  Cypripedium  the  embryo-sac  is 
formed  from  two  megaspores,  which  are  not  separated  by  walls.  In 
Peperomia  the  megaspores  are  not  separated  by  walls,  and  each 
megaspore  nucleus  divides  twice,  so  that  a  16-nucleate  sac  is  formed. 

The  reduction  of  chromosomes  takes  place  during  the  two  mitoses 
by  which  the  mother  cell  gives  rise  to  four  megaspores.  The  figures 
are  much  larger  than  in  the  corresponding  mitoses  in  spermatogenesis, 
but  so  much  more  tedious  to  secure  that  most  studies  in  reduction 
have  been  based  upon  divisions  in  the  pollen  mother  cell.  Lilium  is 
quite  favorable  for  a  study  of  oogenesis,  but  it  must  be  remembered 
that  it  is  exceptional  in  having  an  embryo-sac  formed  from  four 
megaspores. 

In  very  young  stages,  before  the  appearance  of  the  integument, 
the  ovary  may  be  removed  from  the  flower  and  placed  directly  in  the 
fixing  agent,  but  in  later  stages,  such  as  are  shown  in  Fig.  100,  strips 
should  be  cut  off  from  the  sides  of  the  ovary  in  order  to  secure  more 
rapid  fixing  and  more  perfect  infiltration  with  paraffin.  The  dotted 
lines  in  Fig.  99,  C,  show  about  how  much  should  be  cut  off.  This  is 
a  much  better  plan  than  to  secure  rapid  fixing  and  infiltration  by 
cutting  the  ovary  into  short  pieces,  because  the  ovules  will  be  in 
about  the  same  stage  of  development  throughout  the  ovary,  and  when 
one  finds  desirable  stages  like  those  from  which  these  photomicro- 
graphs were  taken,  it  is  gratifying  to  have  these  pieces  as  long  as 
possible. 

Chromo-acetic  acid,  with  the  addition  of  a  little  osmic  acid,  is 
good  for  fixing  the  entire  series.  Iron-haematoxylin,  with  a  light 
touch  of  orange,  is  best  for  the  chromatin.  For  general  beauty 
and  for  the  achromatic  structures,  the  safranin,  gentian- violet, 
orange  combination  has  not  been  excelled.  The  photomicrographs 
(Figs.  96-98)  illustrating  the  series  from  the  archesporial  cell  (which, 
in  this  case,  is  also  the  primary  sporogenous  cell  and  the  megaspore 
mother  cell)  to  the  four  megaspore  nuclei  will  repay  a  careful  study. 


276 


Methods  in  Plant  Histology 


One  more  mitosis  produces  the  8-nucleate  embryo-sac,  but  Lilium 
is  not  a  good  type  for  illustrative  purposes,  since  the  egg  apparatus 
is  not  very  definitely  organized. 

For  the  embryo-sac  at  the  fertilization  stage,  many  of  the  Com- 
positae  are  good.  Senedo  aureus  is  quite  favorable,  because  it  is 
easy  to  cut  and  the  akenes  do  not  spread.  Aster  gives  an  exceptional 

view  of  the  antipodal  re- 
gion, but  is  rather  hard  to 
cut.  Before  fixing,  trim  the 
head  as  indicated  in  Fig.  99. 
Silphium,  especially  S.  lacini- 
atum,  furnishes  an  ideal  view 
of  the  embryo-sac.  With 
thumbs  and  fingers  grasp 
the  two  wings  of  the  akene 
and  carefully  split  it,  expos- 
ing the  single  white  ovule  in- 
side. This  is  rather  tedious, 
but  every  ovule  will  yield  a 
perfectly  median  longitudinal 
section  of  the  embryo-sac, 
and  there  is  not  the  slightest 
difficulty  in  cutting.  When 
the  rays  look  their  best,  the 
embryo-sac  is  ready  for  fer- 
tilization, or  the  pollen  tubes 
may  be  entering;  as  the  rays 
begin  to  wither,  you  will  find  fertilization  or  early  stages  in  the 
embryo  and  endosperm.  Sections  should  be  about  10  p  thick. 

The  Ranunculaceae,  especially  Anemone  patens  var.   Wolfgan- 

giana,  show  a  rather  large,  broad  embryo-sac,  with  highly  organized 

egg  apparatus  and  antipodals.     Sections  should  be  10  to  20  fjt,  thick. 

For  general  views  of  the  embryo-sac,  the  safranin,  gentian-violet, 

orange  combination  is  recommended. 

Fertilization. — The  later  stages  cut  to  show  the  mature  embryo- 
sac  will  often  show  fertilization.  The  male  and  female  nuclei  almost 


FIG.  99. — A,  head  of  Aster;  B,  pod  of  Cap- 
sella;  C,  transverse  section  of  ovary  of  Lilium. 
The  dotted  lines  show  how  the  material  should 
be  trimmed  before  fixing. 


Spermatophytes — Angiosperms  277 

invariably  show  a  difference  in  staining  capacity  when  the  male 
nuclei  are  just  discharged  from  the  pollen  tube.  With  cyanin  and 
erythrosin,  the  male  nucleus  stains  blue  and  the  female  red;  hence 
the  obsolete  terms  cyanophilous  and  erythrophilous.  As  the  nuclei 
come  into  contact  within  the  egg,  they  begin  to  stain  alike,  the  male 
nucleus  staining  more  and  more  like  the  female.  In  the  final  stages 
of  fusion  it  is  difficult,  or 
impossible,  to  distinguish 
the  two  nuclei.  The  male 
nucleus  which  takes  part  in 
the  "triple  fusion"  to  form 
the  endosperm  nucleus  be- 
haves in  the  same  way. 

Lilium  is  a  very  good 
and  always  available  type 
for  illustrating  fertilization 
(Fig.  100).  Take  ovaries 
from  flowers  whose  petals 
have  withered  but  have  not 
yet  fallen  off.  Though 
much  smaller,  Silphium  is 
a  good  type,  because  its 
curved  or  twisted  male  nu- 
clei are  easily  distinguished 
from  the  spherical  nuclei 
in  the  embryo-sac.  The 
embryo-sacs  of  orchids  are 
very  small,  but  ovules  are 
extremely  numerous  and  the  chances  for  securing  the  fusion  of  nuclei 
are  correspondingly  good.  In  Cypripedium  the  nuclei  do  not  fuse  in 
the  resting  condition,  but  the  chromosomes  of  the  two  parents  are 
perfectly  distinct  in  the  egg.  The  general  statement  is  that  nuclei 
fuse  in  the  "resting  condition." 

The  Endosperm. — Some  of  the  preparations  intended  for  fertili- 
zation will  be  likely  to  show  early  stages  in  the  development  of 
endosperm. 


FIG.  100. — Lilium  philadelphicum:  photomi- 
crograph of  section  showing  fertilization  and 
also  the  triple  fusion;  from  a  preparation  and 
negative  by  Dr.  W.  J.  G.  Land.  X585. 


278 


Methods  in  Plant  Histology 


In  rather  long,  narrow  embryo-sacs,  a  cell  wall  is  likely  to  follow 
even  the  first  division  of  the  endosperm  nucleus,  so  that  the  endo- 
sperm is  cellular  from  the  beginning.  Ceratophyllum,  Monotropa, 
and  Verbena  will  furnish  material  of  this  type. 

In  large,  broad  embryo-sacs,  the  formation  of  endosperm  is 
almost  sure  to  be  initiated  by  a  series  of  simultaneous  free  nuclear 


FIG.  101. — Capsella  bursa-pastoris:  A,  first  division  of  the  embryo  cell;  B,  quad- 
'  rants;  C,  octants;  D,  dermatogen  has  been  cut  off;  E,  differentiation  into  periblem  and 
plerome  of  the  root  (the  plerome  cells  are  shaded) ;  F,  the  periblem  of  the  root  is  completed 
at  the  expense  of  the  upper  cell  of  the  suspensor;  G,  the  mitotic  figure  in  the  suspensor 
cell  indicates  that  the  upper  suspensor  cell  by  a  second  contribution  is  about  to  complete 
the  dermatogen  of  the  root;  H,  plerome  (shaded),  periblem,  dermatogen  (shaded),  and 
the  first  layer  of  the  root  cap;  fixed  in  chromo-acetic  acid  and  stained  in  Delafleld's 
haematoxylin;  10  n  thick.  X400. 

divisions.  In  large  sacs  walls  then  begin  to  appear  at  the  periphery 
and  wall  formation  gradually  advances  toward  the  center  until  the 
entire  sac  is  filled  with  tissue.  Lilium,  Peperomia,  and  Ranunculus 
furnish  examples  of  this  type. 


Spermatophytes — A  ngiosperms  279 

An  intermediate  condition  is  seen  in  somewhat  elongated  embryo- 
sacs  of  medium  size,  like  those  of  Compositae.  After  a  few  free 
nuclear  divisions,  walls  appear  simultaneously  throughout  the  entire 
sac.  Silphium  laciniatum  is  particularly  good.  Akenes  from  which 
the  corolla  has  just  fallen  will  furnish  material. 

The  Embryo. — The  common  Capsella  bursa-pastoris  (Shepherd's 
Purse)  is  a  favorable  form  for  a  study  of  the  development  of  a  dicotyl 
embryo.  The  stages  shown  in  Fig.  101,  A-F,  will  be  found  in  pods 
about  3  mm.  in  length.  These  may  be  put  directly  into  the  fixing 
agent,  but  stages  like  G  and  H,  which  are  found  in  pods  about  5  mm. 
in  length,  should  be  trimmed  as  indicated  in  Fig.  99,  B,  before  fixing. 
Formalin  alcohol  is  a  satisfactory  fixing  agent.  Cut  sections  5  to 
10  ju  thick  and  parallel  to  the  flat  face  of  the  pod.  Delafield's  haema- 
toxylin,  without  any  contrast  stain,  is  excellent. 

For  a  study  of  the  monocotyl  embryo,  7ns,  and  especially 
7.  pseudacorus,  can  be  recommended.  The  embryo  is  straight  and 
cotyledon,  stem-tip,  and  root  are  clearly  differentiated  before  the 
endosperm  becomes  too  hard  to  cut  in  paraffin.  Fix  pieces  about 
3  mm.  wide  cut  perpendicular  to  the  face  of  the  cheese-shaped  seed. 
Do  not  try  to  cut  the  whole  pod. 

Sagittaria  has  been  used  quite  extensively.  It  is  easily  obtained, 
the  whole  head  can  be  cut  with  ease,  even  after  the  cotyledon  and 
stem-tip  are  clearly  differentiated,  and  the  endosperm  is  instructive; 
but  the  embryo  is  curved,  like  that  of  Capsella,  and  good  views  are 
rather  rare. 

Zea  Mays,  especially  the  sweet  corn,  is  a  good  type  to  illustrate 
the  peculiar  embryo  of  the  grasses.  Directions  have  been  given  on 
p.  264. 

In  many  forms  good  preparations  of  late  stages  may  b 
by  soaking  the  seeds  in  water  until  the  embryo  bursts  the  seed  coat. 
Young  seedlings  furnish  valuable  material  for  a  study  of  vascular 
anatomy. 


CHAPTER  XXVI 
USING  THE  MICROSCOPE 

The  investigator  who  desires  to  see  all  that  his  microscope  is 
capable  of  showing  must  study  the  optics  of  his  instrument.  The 
fundamental  principles  are  presented  in  any  good  textbook  of  physics. 
Excellent  practical  hints  are  given  in  two  booklets  published  by  the 
leading  American  optical  companies.  These  booklets  tell  the  begin- 
ner how  to  set  up  the  microscope,  how  to  keep  it  in  order,  and  give 
directions  concerning  illumination,  dry  and  immersion  objectives, 
mirror,  condenser,  diaphragm,  and  various  other  things  (Fig.  102). 
They  were  doubtless  written  for  advertising  purposes,  but  since  they 
advertise  by  giving  directions  for  securing  the  best  results  with  the 
microscope,  the  information  is  very  reliable.  The  Spencer  Lens 
Co.,  of  Buffalo,  New  York,  and  The  Bausch  &  Lomb  Optical  Co., 
of  Rochester,  New  York,  furnish  these  booklets  free  of  charge. 

In  the  histological  laboratory  where  preparations  are  being  made 
the  microscope  is  in  constant  danger.  A  cheap  microscope  with  a 
16  mm.  objective  and  one  ocular,  such  an  instrument  as  can  be  got 
for  $20  or  less,  can  be  used  for  examining  preparations  while  they  are 
wet  with  alcohols,  oils,  or  other  reagents.  If  it  is  necessary  to  use 
a  better  instrument  for  such  work,  cover  the  stage  with  a  piece  of 
glass — an  old  lantern  slide  is  of  about  the  right  size — and  be  extremely 
careful  not  to  get  reagents  upon  the  brass  portions. 

MICROMETRY 

Everyone  who  expects  to  become  at  all  proficient  in  the  use  of 
the  microscope  should  learn  to  measure  microscopic  objects  and 
should  learn  to  form  some  estimate  even  without  measuring,  just 
as  one  guesses  at  the  size  of  larger  objects.  In  any  measurement 
one  should  note  the  tube  length,  which  is  usually  160  mm.  Since 
the  use  of  the  nosepiece  is  universal,  it  is  convenient  to  have  the 
length  measure  160  mm.  when  the  tube  is  pushed  in.  Some  com- 
panies still  make  the  tube  so  short  that  it  must  be  pulled  out  about 

280 


Using  the  Microscope 


281 


15  mm.  to  reach  the  length  of  160  mm.,  even  when  the  nosepiece  is 
in  place.  Where  there  is  no  revolving  nosepiece,  the  draw-tube 
is  simply  pulled  out  until  the  length  is  160mm.  (Fig.  103).  Where 
'"?*—-—.. 

OCUCAR - 

—DRAW- TUBE 


OBJECTIVES I_r 


PINION 
COARSE  ADJUSTMENT. 


GRADUATED  SHORT  SLICE 

STAGE-- 
ADJUSTABLE 

SPRING    FINGER 

CONDENSER   MOUNTING 
DROP  -SWING  ARM.--"" 
LOWER  IRIS  DIAPHRAGM-' 
FOR  OBLIOUE  LIGHT. 

STAGE  CENTERING  SCREWS 


RROR  FORK 
MIRROR 

RACK  a  PINION 

BUTTON 


HORSESHOE 
--£>ASE 


FIG.  102.— A  modern  microscope 


a  nosepiece  is  used,  its  height  should  be  measured,  and  the  draw-tube 
should  be  pushed  in  a  distance  equal  to  the  length  of  the  nosep.eee. 
There  are  in  general  use  two  practical  methods  of  measuring  m,cr 
scopic  objects,  one  by  means  of  the  ocular  micrometer,  and  1 
by  means  of  camera  lucida  sketches. 


282 


Methods  in  Plant  Histology 


Measuring  with  the  Ocular  Micrometer. — A  stage  micrometer 
and  an  ocular  micrometer  are  necessary.  A  stage  micrometer  should 
be  ruled  in  tenths  and  one-hundredths  of  a  millimeter.  It  does  not 
matter  what  the  spacing  in  the  ocular  micrometer  may  be,  except 

that  the  lines  must  be  at  equal 
distances  from  each  other.  As  a 
matter  of  fact,  the  ocular  mi- 
crometer is  generally  ruled  in 
tenths  of  a  millimeter,  but  this 
ruling  is  more  or  less  magnified  by 
the  lens  of  the  ocular. 

Place  the  stage  micrometer  up- 
on the  stage  and  the  ocular  mi- 
crometer in  the 'tube,  and  arrange 
the  two  sets  of  rulings  so  that  the 
first  line  in  the  ocular  micrometer 
will  coincide  with  the  first  line  of 
the  stage  micrometer,  and  then 
find  the  value  of  one  space  in  the 
ocular  micrometer.  The  method 
of  finding  this  value  is  shown  in 
the  following  case  in  which  the 
tube  length  was  160  mm.,  the 
ocular  a  Zeiss  ocular  micrometer  2, 
and  the  obj ective  a  Leitz  3.  In  the 
ocular  micrometer,  ninety-eight 
spaces  covered  just  fifteen  of  the 
larger  spaces  of  the  stage  microm- 
eter. Since  the  stage  micrometer 
is  ruled  in  tenths  and  one- 
hundredths  of  a  millimeter,  the 

fifteen  spaces  equal  1.5mm.,  or  1,500 /it.1  Then  98  spaces  of  the 
ocular  micrometer  equal  1,500  ju;  and  one  space  in  the  ocular  equals 
-£$  of  1,500  fj,,  or  15.3  ju.  This  value  being  determined,  there  is  no 
further  use  for  the  stage  micrometer.  To  measure  the  diameter  of 

1  One  millimeter  =  1,000  /a.    The  Greek  letter  ^  is  an  abbreviation  for  M^PO",  or  micron. 


=ps 

|_ 

~) 

I  oftheOcu 

'Diaphracjrr/ 

1 

•^  — 

>                    1     ' 
-•Fieldlens' 

—  DrawTube 

d 

It) 

X 

< 

..-Body  Tub* 

,i 

"c 
O 

_ 

1 

^DrawTube  Diaphragm 
with  Society  Screw 

Ws— 

r^W 

o/lhe 
Objective 


Fio.  103.— Tube  length 


Using  the  Microscope  283 

a  pollen  grain  put  the  preparation  on  the  stage,  using  the  same 
objective  and  ocular  micrometer,  and  note  how  many  spaces  a  pollen 
grain  covers.  If  the  pollen  grain  covers  five  spaces,  its  diameter  is 
five  times  15 . 3  /*,  or  76 . 5  /*.  In  the  same  way,  the  value  of  a  space 
in  the  ocular  when  used  with  the  other  objectives  should  be  de- 
termined. The  values  for  three  or  four  objectives  may  be  written 
upon  an  ordinary  slide  label  and  pasted  upon  the  base  of  the  micro- 
scope for  convenient  reference. 


SsA  ..PlAt\otct,be,QjLuW;la. 


nb  \jcm\sA. 

I/O  rfKcf  misiw  4^  60 


H  bvm- Oho. Obj-  iSi 
yey  "liBfc 

10  30  v»     \  6d  -K    to  ye       //»  ru>  i3»  no 
I     I     I          111!          II     I     I 


FIG.  104. — Scale  card  for  practical  use.  The  figure  is  considerably  smaller  than  the 
card  from  which  it  was  made. 

This  method  is  the  best  one  for  measuring  spores  and  for  most 
measurements  in  taxonomy. 

Measuring  by  Means  of  Camera  Lucida  Sketches.— This 
method  is  of  great  importance  in  research  work,  because  vari- 
ous details  can  be  measured  with  far  greater  rapidity  than 
by  the  other  method.  Upon  a  piece  of  cardboard,  about  as 
thick  as  a  postal  card,  draw  a  series  of  scales  like  those  shown  in 
Fig.  104. 

Make  a  scale  for  each  objective.  It  is  not  necessary  to  make 
scales  for  all  the  oculars,  but  only  for  the  one  in  most  constant  use. 


284  Methods  in  Plant  Histology 

It  is  absolutely  necessary  to  note  the  tube  length,  length  of  the  bar 
of  the  camera  mirror  and  inclination  of  the  camera  mirror,  and  the 
level  at  which  the  scale  is  made.  A  variation  in  any  of  these  details 
will  change  the  scale. 

In  using  the  stage  micrometer,  place  the  cardboard  on  the  table, 
and  with  the  aid  of  the  camera  lucida  sketch  the  rulings  of  the  microm- 
eter. In  Fig.  104  (which  has  been  reduced  by  photography)  note, 
for  example,  the  scale  drawn  for  Leitz  objective  3.  The  spaces  are 
drawn  from  the  tenths  of  a  millimeter  rulings  of  the  stage  microm- 
eter. Therefore  each  space  on  the  card  represents  one-tenth  of  a 
millimeter,  or  100  /*,  and  the  ten  spaces  shown  on  the  card  represent 
1  mm.,  or  1,000  p.  By  measuring  with  a  metric  rule  the  ten  spaces 
upon  the  card,  it  is  found  that  the  scale  is  114  mm.  in  length.  The 
magnification  of  any  drawing  made  with  the  same  ocular  and  objec- 
tive, under  the  same  conditions,  will  therefore  be  114  diameters. 
This  does  not  mean  that  the  magnifying  power  of  Leitz  objective  3 
with  Zeiss  ocular  4  is  114  diameters,  for  the  magnification  of  this 
combination  is  much  less.  A  scale  drawn  at  the  level  of  the  stage 
would  show  more  nearly  the  magnifying  power  of  the  combination, 
but  would  still  give  too  large  a  figure.  The  exact  size  of  any  object 
which  has  been  sketched  with  this  combination  can  now  be  measured 
by  applying  the  cardboard  scale,  just  as  one  would  measure  gross 
objects  with  a  rule. 

The  diameter  of  the  field  with  this  combination  is  1,800  /z.  By 
knowing  the  diameter  of  the  field  with  the  various  combinations,  one 
can  guess  approximately  the  size  of  objects. 

Other  combinations  are  made  in  the  same  way.  An  ex- 
cellent check  on  the  accuracy  of  the  computations  is  to  measure 
the  same  object  by  means  of  the  ocular  micrometer  and  by  the 
scale  card.  If  the  results  are  the  same,  the  computations  are 
correct. 

In  making  sketches,  it  is  a  good  plan  to  add  the  data  which  would 
be  needed  at  any  time  in  making  measurements;  e.g.,  L.  3,  Z.  oc.  4, 
table,  110,  50°,  would  show  that  the  sketch  was  made  with  Leitz 
objective  3,  Zeiss  ocular  4,  at  the  level  of  the  table,  with  mirror  bar 
at  110,  and  camera  mirror  at  50°. 


Using  the  Microscope  285 

ARTIFICIAL  LIGHT 

During  a  considerable  part  of  the  year  daylight  is  often  insufficient 
for  successful  work  with  the  microscope.  Numerous  contrivances 
for  artificial  illumination  have  been  devised,  some  of  them  fairly  good, 
but  most  of  them  thoroughly  unsatisfactory.  More  than  two  hun- 
dred years  ago  Hooke  used  a  device  for  artificial  illumination  which 
probably  suggested  the  apparatus  used  by  the  late  Professor  Stras- 
burger  at  Bonn.  The  apparatus  in  use  in  our  own  laboratory  is 
only  a  slightly  modified  form  of  that  used  in  the  Bonn  laboratory. 

The  apparatus  consists,  essentially,  of  a  hollow  sphere  filled  with 
liquid.  A  fairly  good  and  practical  light  can  be  got  with  an  ordinary 
lamp  by  allowing  the  light  to  pass  through  a  wash  bottle  filled  with  a 
weak  solution  of  ammonia  copper  sulphate.  A  piece  of  dark  paper 
with  a  circular  hole  in  it  serves  as  a  diaphragm,  and  at  the  same  time 
protects  the  eyes  from  the  direct  light  of  the  lamp.  Such  an  arrange- 
ment is  shown  in  Fig.  105.  Wash  bottles,  however,  are  not  perfectly 
spherical  and  the  mounting  is  not  convenient.  To  secure  a  perfectly 
spherical  globe,  it  was  necessary  to  have  a  mold  made.  The  globes, 
as  we  now  use  them,  are  of  the  finest  flint  glass,  have  a- diameter  of 
15cm.  (6  inches),  and  are  mounted  in  a  convenient  black  frame, 
Fig.  106.  The  globe  acts  not  only  as  a  condenser,  but  also  as  a  ray 
filter.  For  general  laboratory  work  and  for  nearly  all  research  work, 
a  weak  solution  of  ammonia  copper  sulphate  has  proved  most  satis- 
factory. The  solution  (to  fill  one  15  cm.  globe)  may  be  made  by 
adding  50  c.c.  of  ammonia  to  25  c.c.  of  a  10  per  cent  solution  of  copper 
sulphate,  and  then  adding  enough  distilled  water  to  fill  the  globe. 
If  a  white  precipitate  appears  and  makes  the  solution  look  milky, 
add  more  ammonia.  The  strength  of  the  solution  depends  so  much 
upon  the  power  of  the  light  that  no  fixed  formula  can  be  given. 
Simply  dissolve  in  water  a  small  crystal  of  copper  sulphate— about 
as  large  as  a  grain  of  corn— then  add  about  50  c.c.  of  ammonia,  and 
then  add  distilled  water  until  a  light,  clear-blue  solution  is  secured. 
With  a  very  strong  light,  the  solution  may  have  a  rather  deep-blue 
color;  with  a  less  powerful  light,  the  solution  must  be  weaker. 

In  studying  the  extremely  difficult  achromatic  structures  con- 
cerned in  nuclear  division,  a  light  violet  solution  of  permanganate 


286  Methods  in  Plant  Histology 

of  potash  is  a  good  filter,  if  the  preparation  has  been  stained  in  violet. 
Similarly,  various  niters  may  be  used  according  to  the  staining  of  the 
more  critical  structures. 

The  Welsbach  lamp  furnishes  an  excellent  light.  It  should  be 
placed  so  that  the  rays  will  be  focused  upon  the  mirror  of  the  micro- 
scope. Some  of  the  more  powerful  acetylene  bicycle  lamps  are  quite 


FIG.  105. — A  wash  bottle  used  as  a  condenser 

satisfactory.  The  Argand  type  of  gaslight  is  good,  but  will  usually 
need  a  reflector  behind  it.  A  kerosene  lamp  must  also  be  reinforced 
by  a  reflector.  The  old-fashioned  silvered  reflector,  still  used  in 
country  churches  and  halls,  will  do,  but  is  hardly  equal  to  the  cheap 
reflectors  of  shorter  focus  which  are  so  commonly  used  with  incandes- 
cent electric  lights.  The  incandescent  electric  light  itself  has  not 
given  satisfactory  results.  Small  electric  arc  lights,  tempered  with 
ground  glass  and  filters,  are  very  satisfactory;  the  only  objection  is 


Using  the  Microscope  287 

that  they  require  frequent  adjustment.     The  Nernst  light  is  £ 
source  of  illumination,  but,  after  repeated  trials,  we  have  laid  it 
aside  because  it  is  always  getting  out  of  order. 

Optical  companies  are  now  making  very  small  incandescents  with 
reflector,  ground  glass,  and  filter  which  promise  to  be  satisfactory 
for  most  work. 


Fir;.    lOfi.  —  A  satisfactory  artificial  light 


We  are  still  using  the  globes  and  find  such  light  not  only  equal 
to  the  best  daylight,  but  in  many  cases  superior. 

When  using  the  camera  lucida,  it  is  necessary  to  have  a  mirror 
placed  so  as  to  throw  a  fairly  strong  light  upon  the  paper  and  the 
pencil  point.  A  piece  of  silvered  glass  3  or  4  inches  square  is  large 
enough.  Such  a  mirror  can  be  held  by  an  ordinary  ring  stand,  as 
shown  in  Figs.  105  and  106. 


CHAPTER  XXVII 

LABELING  AND  CATALOGING  PREPARATIONS 
THE  LABEL 

We  should  say  that  the  first  thing  to  write  upon  a  label  is  the 
genus  and  species  of  the  plant;  the  next  thing  would  be  the  name  of 
the  organ  or  tissue,  and  then  might  be  added  the  date  of  collection; 
e.g.,  Marchantia  polymorpha,  young  archegonia,  January  10,  1915. 
The  date  of  making  the  preparation  is  of  no  value  unless  the  student 
is  testing  the  permanence  of  stains  or  something  of  that  sort.  It 
is  hardly  worth  while  to  write  upon  the  label  the  names  of  the  stains 
used,  for  the  student  will  soon  learn  to  recognize  the  principal  stains. 
A  hasty  sketch  on  the  label  will  often  indicate  any  exceptionally 
interesting  feature  in  the  preparation.  To  facilitate  finding  such  a 
feature,  it  is  a  good  plan  to  mark  the  particular  section  or  sections  with 
ink,  the  marking  being  always  on  the  under  side  of  the  slide  so  as 
not  to  cause  any  inconvenience  if  an  immersion  lens  should  be  used. 

CATALOGING  PREPARATIONS 

As  a  collection  grows,  the  student  will  need  some  device  for 
locating  readily  any  particular  preparation.  Some  have  their  slides 
numbered  and  cataloged,  but  all  devices  of  this  sort  are  too  cumbrous 
and  slow  for  the  practical  worker  in  the  laboratory.  After  twenty 
years'  experience  with  a  collection  which  now  numbers  more  than 
twenty-five  thousand  preparations,  we  recommend  the  following 
system : 

Four  wooden  slide  boxes  of  the  usual  type  will  do  for  a  beginning; 
they  should  be  labeled:  THALLOPHYTES,  BRYOPHYTES,  PTERIDO- 
PHYTES,  and  SPERMATOPHYTES.  As  the  collection  grows  and  new 
boxes  are  needed,  the  classification  can  be  made  more  definite;  e.g., 
there  should  be  a  box  labeled  BRYOPHYTES  Hepaticae  and  one  labeled 
BRYOPHYTES  Musd.  As  the  liverwort  collection  grows,  three  boxes 
will  be  necessary,  and  should  be  labeled  BRYOPHYTES  Hepaticae 

288 


Labeling  and  Cataloging  Preparations 


289 


Marchantiales,  BRYOPHYTES  Hepaticae  Jungermanniales,  and  BRYO- 
PHYTES  Hepaticae  Anthocerotales.  It  will  readily  be -seen  that  the 
process  can  be  continued  almost  indefinitely,  and  that  new  slides 
may  be  at  any  time  dropped  into  their  proper  places.  A  rather 
complete  label  gradually  built  up  in  this  way  is  shown  in  Fig.  107: 


BRYOPHYTES 

HEPATICAE 

Ju  nge  rm  a.  n  n  ia  les 
Porella    platyphyllum 


Archegonia 


FIG.   107. — A  label  for  slide  boxes 

The  beginner  will  often  find  that  the  mere  placing  of  a  slide  in  the 
proper  box  and  the  box  in  its  proper  place  on  the  shelf  will  refresh  or 
increase  his  knowledge  of  classification.  While  this  system  is  almost 
ideal  for  the  careful  worker,  especially  if  he  has  some  knowledge  of 
classification,  it  is  the  worst  possible  method  for  a  careless  student, 
since  a  slide  in  the  wrong  box  is  almost  hopelessly  lost  if  the  collec- 
tion is  large  enough  to  need  thorough  classifying. 


CHAPTER  XXVIII 
A  CLASS  LIST  OF  PREPARATIONS 

Where  a  regular  course  in  histology  is  conducted,  it  is  a  good  plan 
to  give  each  student  at  the  outset  a  complete  list  of  the  preparations 
which  he  is  expected  to  make.  In  a  three  months'  course  a  fairly 
representative  collection  of  preparations  can  be  made.  The  avail- 
ability of  material  determines  what  a  list  shall  be.  Besides  gaining 
an  introduction  to  the  use  of  the  microscope  and  its  accessories,  a  class 
meeting  ten  hours  a  week  for  twelve  weeks  should  be  able  to  do  as 
much  work  as  is  outlined  below. 

In  making  the  mounts  the  order  indicated  in  the  list  should  not 
be  followed.  Begin  with  temporary  mounts,  -and  then  study,  in 
succession,  freehand  sections  (the  glycerin  method),  the  Venetian 
turpentine  method,  the  paraffin  method  (the  celloidin  method),  and 
special  methods.  A  large  proportion  of  the  time  should  be  devoted 
to  the  paraffin  method. 

It  is  neither  possible  nor  desirable  that  each  student  should  in 
every  case  go  through  all  the  processes  from  collecting  material  to 
labeling.  Some  of  the  material  may  be  in  85  per  cent  alcohol,  some 
in  formalin,  some  in  glycerin,  some  in  Venetian  turpentine,  and  some 
in  paraffin.  One  student  may  imbed  in  paraffin  enough  of  the 
Anemone  for  the  whole  class;  another  may  imbed  the  Lilium  stamens; 
and  by  such  a  division  of  labor  a  great  variety  of  preparations  may 
be  secured  without  a  corresponding  demand  upon  the  time  of  the 
individual. 

LIST  OF  PREPARATIONS 

THALLOPHYTES 
SCHIZOPHYTES 

MYXOMYCETES 

1.  Trichia  varia. — Paraffin  sections,  5/x.    Safranin,  gentian- violet,  orange. 

SCHIZOMYCETES 

2.  Bacteria. — Coccus,  Bacillus,  and  Spirillum  forms.    Stain  on  cover-glass 
or  slide. 

3.  Bacillus  anthracis.—ln  liver  of  mouse.    Paraffin  sections,  5/x.    Stain 
in  gentian- violet,  Gram's  method. 

290 


A  Class  List  of  Preparations  291 

CYAXOPHYCEAE  (SCHIZOMYCETES) 

4.  OsciUatoria.-Put  living  material  into  10  per  cent  glycerin  and  allow 
it  to  concentrate. 

5.  Tolypothrix.—Use    the    Venetian    turpentine    method.    Should    show 
heterocysts,  hormogonia,  and  false  branching. 

6.  Nostoc.— Venetian  turpentine  method. 

7.  Wasserbluthe.—The  principal  forms  in  this  material  are: 

a)  Coelosphaerium   Kutzingianum.— Colonies   in    the   form   of   hollow 
spheres. 

b)  Anabaena  gigantea. -Filaments  straight.     Preparations  should  show 
vegetative  cells,  heterocysts,  hormogonia,  and  spores. 

c)  Anabaena  flos-aquae.— Filaments  curved.     Stain  on  the  slide  and 
mount  in  balsam. 

8.  Gloeotrichia— Smear  on  the  slide,  stain  in  safranin  and  gentian- violet, 
and  mount  in  balsam;   or  use  the  Venetian  turpentine  method,  staining 
in  Magdala  red  and  anilin  blue  and  crushing  under  the  cover-glass. 

ALGAE 
CHLOROPHYCEAE 

9.  Volvox.—Vse  the  Venetian  turpentine  method.     If  paraffin  material 
is  available,  cut  5  /*  in  thickness  and  stain  in  safranin,  gentian-violet, 
orange. 

10.  Scenedesmus. — Let  a  drop  containing  the  material  dry  upon  the  slide, 
stain,  and  mount  in  balsam. 

11.  Hydrodictyon. — Use  the  Venetian  turpentine  method. 

Each  preparation  should  contain  pieces  of  old  and  of  young  nets,  and 
also  at  least  one  young  net  developing  within  an  older  segment.  The  great- 
est care  must  be  taken  not  to  injure  the  older  segments  while  arranging  the 
mount. 

12.  Ulothrix. — Use  the  Venetian  turpentine  method.     Each  mount  should 
show  various  stages  in  the  development  of  spores  and  gametes. 

13.  Oedogonium. — Stain  in  Magdala  red  and  anilin  blue  and  mount  in 
Venetian  turpentine. 

14.  Coleochaete. — Stain  in  Delafield's  haematoxylin  and  mount  in  balsam. 

15.  Cladophora. — Stain  some  in  iron-haematoxylin  and  some  in  Magdala 
red  and  anilin  blue.     Mount  both  together  in  Venetian  turpentine. 

16.  Diatoms.— 'Make  mounts  of  the  frustules  and  also  stained  preparations 
showing  the  cell  contents. 

17.  Desmids—  Make  mounts  of  available  forms.     Use  the  Venetian  turpen- 
tine method  if  material  is  sufficiently  abundant. 

18.  Zygnema.—St&m  in  iron-haematoxylin  and  mount  in  Venetian  turpen- 
tine. 


292  Methods  in  Plant  Histology 

19.  Spirogyra. — Stain  in  Magdala  red  and  anilin  blue,  and  mount  in  Venetian 
turpentine. 

20.  Vaucheria. — Stain  in  Magdala  red  and  anilin  blue,  and  mount  in  Venetian 
turpentine. 

21.  Chara. — Cut  paraffin  sections  of  the  apical  cell,  oogonia,  and  antheridia. 

PHAEOPHYCEAE 

22.  Ectocarpus. — Stain  some  in  iron-haematoxylin  and  some  in  Magdala 
red  and  anilin  blue.    Mount  both  together  in  Venetian  turpentine. 

23.  Cutleria. — Sections  of  oogonia,  antheridia,  and  sporangia.    Cut  10  p 
thick  and  stain  in  iron-haematoxylin  with  about  7  minutes  in  safranin. 

24.  Fucus    vesiculosus. — Antheridial    conceptacle    with    paraphyses    and 
antheridia;    oogonial  conceptacle  with  oogonia.    Cut  10 /u,  thick  and 
stain  in  iron-haematoxylin  with  about  5  minutes  in  safranin. 

KHODOPHYCEAE 

25.  Nemalion. — Stain  some  in  iron-haematoxylin  and  -some  in  eosin.    Each 
preparation  should  show  trichogyne,  carpogonium,  and  cystocarp.    You 
cannot  mount  it  in  Venetian  turpentine:  use  glycerin  or  glycerin  jelly. 

26.  Polysiphonia. — Stain  in  iron-haematoxylin  or  Magdala  red  and  anilin 
blue.    Mount   whole   in   Venetian   turpentine.    Each   mount   should 
show  tetraspores,  antheridia,  and  cystocarps.     If  material  is  in  paraffin, 
cut  sections  about  7  ft  thick. 

FUNGI 
PHYCOMYCETES 

27.  Mucor  stolonifer. — Stain  young  sporangia  in  eosin,  dilute  Delafield's 
haematoxylin,  or  in  Magdala  red  and  anilin  blue.     Zygosporic  material 
may  be  mounted  without  staining  or  after  a  very  light  staining  in  dilute 
Delafield's  haematoxylin.    Eosin  is  also  good.     Mount  in  Venetian 
turpentine. 

28.  Saprolegnia. — Stain  in  Magdala  red  and  anilin  blue.    Mount  in  Venetian 
turpentine.    Each  mount  should  show  sporangia  and  oogonia. 

29.  Albugo  (Cijstopus)  candi dus. —Select  white  blisters  which  have  not  yet 
broken  open.    Paraffin,  5  p.    Iron-haematoxylin  and  orange. 

30.  Albugo  bliti  on  Amarantus  retroflexus. — Cut  out  small  portions  of  leaves 
in  which  the  oogonia  can  be  seen  in  abundance.    Paraffin,  5  /A. 

ASCOMYCETES 

31.  Peziza. — Paraffin  sections  of  young  apothecia,  5 /u,  or  less;   sections  of 
older  apothecia,  10  or  15  p.    Safranin,  gentian-violet,  orange. 

32.  Aspergillus,  Eurotium. — Stain  in  eosin  and  mount  in  glycerin,  or  stain 
in  Magdala  red  and  anilin  blue,  and  mount  in  Venetian  turpentine. 

33.  Penicillium. — Treat  like  Aspergillus. 


A  Class  List  of  Preparations  293 

34.  Erysipke  commune  on  Polygonum  aviculare. — Strip  the  fungus  from  the 
leaf.     Paraffin,  5  /j.  or  less.     Safranin,  gentian-violet,  orange. 

35.  Uncinula  necator  on  Ampelopsis  quinquefolia. — Stain  in  Magdala  red 
and  anilin  blue.     Mount  whole  in  Venetian  turpentine  and  break  the 
perithecia  under  the  cover. 

36.  Xylaria—  Paraffin  sections  of  younger  stages.     Delafield's  haematoxylin 
and  erythrosin.     Be  sure  that  some  section  in  each  mount  shows  the 
opening  of  a  perithecium. 

LICHENS 

37.  Physcia  stellaris. — Cut  in  paraffin,  5  p.    Stain  in  cyanin  and  erythrosin. 

BASIDIOMYCETES 

38.  Puccinia   graminis. — Aecidium   stage   on   barberry   leaf.     Uredospore 
and  teleutospore  stage  on  oats.     Cut  3  //.  and  stain  in  iron-haematoxylin. 

39.  Coprinus  micaceus. — Paraffin.    Transverse   sections  of    gills    showing 
trama,  paraphyses,  basidia,  and  spores.    To  show  the  basidium  with 
four  spores,  the  sections  should  be  15  p.  thick.     For  development  of  the 
spores,    cut    5  //,   or   less.    Safranin,    gentian-violet,    orange.    Boletus, 
Hydnum,  and  Polijporus  are  treated  in  the  same  manner. 

BRYOPHYTES 

HEPATICAE 

40.  Riccia natans—  Paraffin,  10  or  15 \i.     Delafield's  haematoxylin.     Arche- 
gonia,  antheridia,  and  sporophytes  imbedded  in  the  gametophyte. 

41.  Marchantia  polymorpha.—ParaSai,  5  or  10  p.    Archegonia,  antheridia, 
and  sporophytes. 

42.  Anthoceros  laevis—  Paraffin,  5  or  10 /*.     Longitudinal  and  transverse 
sections  of  sporophyte.     Safranin,  gentian-violet,  orange. 

43.  Pellia  epiphylla.— Paraffin,  5  or  10  /*.    Longitudinal  sections  of  sporo- 
phyte attached  to  gametophyte.     Safranin,  gentian-violet,  orange. 

44.  Porella  platyphyllum.—P&T&ffin,  10  p..    Delafield's  haematoxylin.    Arche- 
gonia, antheridia,  sporophyte,  and  apical  cell. 

MUSCI 

45.  Sphagnum.— Leaf  buds.     Cut  5  /*  and  stain  in  safranin  and  anilin  blue. 

46.  Sphagnum.— Capsule.    Paraffin.     Delafield's  haematoxylin  and  eryth- 
rosin. 

47.  Funaria  hygrometrica.—'P&r&ttiL.    Longitudinal  and  transverse  s< 
of  young  capsules.     Delafield's  haematoxylin. 

48    Funaria  hygrometrica  or  any  favorable  form.     Protonema.     Place  the 
well-cleaned  material  directly  into  10  per  cent  glycerin  and  allow  it 
concentrate.     Mount  in  glycerin  or  glycerin  jelly. 

49.  Bryum  proZtfenm.-Paraffin.    Antheridia,    10/t;    arch 
20  u;  capsule,  10 /*. 


294  Methods  in  Plant  Histology 

PTERIDOPHYTES 

LYCOPODIALES 

50.  Lycopodium  lucidulum— Transverse  section  of  stem.    Freehand  sections. 
Safranin  and  Delafield's  haematoxylin. 

51.  Lycopodium  inundatum. — Paraffin.     Longitudinal  sections  of  strobilus. 

52.  Selaginella. — Paraffin.    Longitudinal  sections  of  rather  mature  strobili. 
Cyanin  and  erythrosin,  or  safranin,  gentian-violet,  orange. 

53.  Isoetes  echinospora.—Tr&nsverse  section  of  stem.    Paraffin.     Safranin 
and  Delafield's  haematoxylin. 

54.  Isoetes  echinospora. — Paraffin.    Longitudinal  sections  of  microsporangia 
and  megasporangia.    Safranin,  gentian- violet,  orange. 

EQUISETALES 

55.  Equisetum  arvense. — Prothallia  in  Venetian  turpentine.    Stem-tips  in 
paraffin.    Transverse  section  of  stem  freehand  or  in  celloidin. 

OPHIOGLOSSALES 

56.  Botrychium  virginianum. — Paraffin.    Stain  rhizome,   stipes,  and  root 
in  safranin  and  Delafield's  haematoxylin.    Stain  sporangia  in  iron- 
haematoxylin. 

FILICALES 

57.  Protostele. — Use  Gleichenia.    Cut  freehand  and  stain  in  safranin  and 
anilin  blue. 

58.  Solenostele  (amphiphloic  siphonostele). — Use  Adiantum. 

59.  Ectophloic  siphonostele. — Use  Osmunda  cinnamomea. 

60.  Polystele. — Use  Pteris  aquilina  or  any  species  of  Polypodium. 

61.  Sporangia. — For  development,  use  Pteris,  Aspidium,   Cyrtomium,  or 
try  any  available  species.    For  mitosis,  Osmunda  is  exceptionally  good. 

62.  Antheridia   arid   archegonia. — Mount   whole   in   Venetian   turpentine. 
Magdala  red  and  anilin  blue.    Sections  should  be  5  to  10  /A  thick. 
Stain  in  iron-haematoxylin  and  orange. 

63.  Embryo. — Pteris  and  Adiantum  are  good.    Cut  longitudinal  vertical 
sections  10  /*  thick. 

SPERMATOPHYTES 

GYMNOSPERMS 

CYCADALES 

64.  Zamia—  Freehand  sections  of  stem.    Safranin  and  light  green.    Trans- 
verse sections  of  microsporophyll,  5  or  10  /x.     Longitudinal  sections  of 
entire  ovule,    10  to   15  /A;    stain  in  safranin,   gentian- violet,   orange. 
Longitudinal    sections    of    nucellus    with    pollen   tubes,    10  //,.    Iron- 
haematoxylin  and  orange. 


A  Class  List  of  Preparations  295 

GINKGOALES 

65.  Ginkgo  biloba. — Longitudinal  sections  of  endosperm  showing  archegonia 
or  young  embryos.     Paraffin  10  //,. 

Sections  of  microsporangia  with  nearly  mature  pollen,  5  /<.. 

CONIFERALES 

66.  Pinus  Laricio. — Transverse  sections  of  needles  and  young  stem.     Free- 
hand or  celloidin.     Safranin  and  Delafield's  haematoxylin. 

67.  Pinus   Slrobus. — Freehand   sections   of   well-seasoned   wood.     Methyl 
green  and  fuchsin,  or  safranin  and  Delafield's  haematoxylin. 

68.  Pinus  Laricio. — Paraffin.     Longitudinal  section  of  mature  staminate 
strobilus.     Safranin,  gentian- violet,  orange. 

69.  Abies  balsamea  or  Pinus  Laricio. — Pollen  at  shedding  stage  shaken  out 
and  imbedded  in  paraffin;  5  //,.     Stain  in  safranin,  gentian-violet,  orange. 

70.  Pinus  Laricio. — Paraffin.     Ovule  with  archegonia.     Safranin,  gentian- 
violet,  orange. 

71.  Pinus    sylvestris    or    P.    Laricio. — Paraffin.     Embryos.     Cyanin    and 
erythrosin,  or  safranin  and  anilin  blue. 

GNETALES 

72.  Transverse  section  of  stem  of  Ephedra.    Freehand. 

73.  Longitudinal  section  of  the  ovule  of  Ephedra. 

ANGIOSPERMS 
DICOTYLS 

74.  Pelargonium.— Transverse   sections   of   stem   to   show   phellogen   and 
intrafascicular  cambium.     Freehand.     Endarch  siphonostele. 

75.  Tilia  americana.— Celloidin  or  freehand.    Transverse  sections  of  small 
stems  3  mm.  to  6  mm.  in  diameter.     Safranin  and  Delafield's  haematoxy- 
lin.    Endarch  siphonostele  with  annual  rings. 

76.  Sambucus  nigra.— Transverse  section  of  primary  root  to  show  origin 
of  secondary  structures. 

77.  Cucurbita—  Longitudinal  section  of  stem  to  show  sieve  tubes. 

78.  Capsella bursa-pastoris—  Paraffin.     Floral  development,  5 p..    Embryos, 
5  to  10  p.     Stain  both  in  Delafield's  haematoxylin  without  any  contrast 
stain. 

79.  Taraxacum  officinale.— Paraffin.     Floral  development,   5  p..    Embryo- 
sac,  10  to  15  //.. 

80.  Ranunculus.— Longitudinal  sections  of  young  flowers  to  show  megaspor 
mother  cells  and  megaspores. 

81    Silphium.— Longitudinal  sections  of  the  ovule  at  the  fertilization  per 
Longitudinal  sections  of  staminate  flowers  just  before  the  s 
pollen. 

82.  Anemone  patens.— Paraffin.    Embryo-sac. 


296  Methods  in  Plant  Histology 

MONOCOTYLS 

83.  Clintonia. — Transverse  section  of  stem  to  show  siphonostele.     Freehand. 
Safranin  and  anilin  blue. 

84.  Acorus  calamus. — Transverse  sections  of  rhizome,  freehand  or  paraffin, 
to  show  amphi vasal  bundles. 

85.  Zea  Mays. — Transverse  section  of  stem  to  show  scattered  bundles;  also 
good  for  companion  cells.    Freehand.    Safranin  and  anilin  blue. 

86.  Tradescantia   virginica. — Longitudinal   sections   of   root-tip.     Paraffin, 
5  and  10  /u,.    Stain  for  mitosis. 

87.  Smilax  herbacea. — Transverse  section  of  adult  root.    Freehand.    Shows 
exarch,  radial  structure,  and  a  highly  developed  endodermis.    Safranin 
and  Delafield's  haematoxylin. 

88.  Lilium. — Transverse  section  of  leaf.    Freehand.    Transverse  sections 
of  ovaries  in  various  stages  from  megaspore  mother  cell  to  fertilizations; 
transverse  sections  of  anthers  to  show  microspore  mother  cells  and  reduc- 
tion of  chromosomes;    also  later  stages  with  nearly  mature  pollen. 
Paraffin  5  to  10  ^. 

89.  Iris. — Section  of  young  seeds   to  show  embryo  with   cotyledon  and 
stem-tip. 

90.  Sagittaria. — Longitudinal  sections  of  ovulate  flowers  of  various  stages 
to  show  development  of  the  embryo  and  endosperm. 

91.  Zea  Mays. — Longitudinal  and  transverse  sections  of  embryo  (sweet  corn, 
roasting-ear  condition)  to  show  structure  of  root  and  beginning  of  pro- 
toxylem. 


CHAPTER  XXIX 
FORMULAE  FOR  REAGENTS 

FIXING  AGENTS 
Absolute  Alcohol.— Used  alone  without  any  mixtures. 

Carnoy's  Fluid.— 

Absolute  alcohol 2  Parts 

Chloroform 3  Parts 

Glacial  acetic  acid •   1  Part 

Farmer's  Fluid. — 

Absolute  alcohol •  6  Parts 

Glacial  acetic  acid •   *  Part 

Formalin  Alcohol  (Lynds  Jones's  formula) .- 

70  per  cent  alcohol 

Commercial  formalin •.  •  • 

Formalin  Alcohol  (Chicago  formula).— 
70  per  cent  alcohol.. 
Commercial  formalin 

Formalin.— 

Commercial  formalin 

,  1UU  C.C. 

Water 

Stock  Chromo-Acetic  Solution.— 

1  K. 
Chromic  acid 1  c'c 

Glacial  acetic  acid .  .  '          c'c' 

Water 

Schaffner's  Chromo-Acetic  Solution .— 

Chromic  acid 0  7  c  c_ 

Glacial  acetic  acid .  .  99  0  c.c. 

Water 

297 


298  Methods  in  Plant  Histology 

Chromo- Acetic  Solution  (for  delicate  structures). — 

Chromic  acid 1  g. 

Glacial  acetic  acid 3  c.c. 

Water 300  c.c. 

The  addition  of  10  drops  of  osmic  acid  to  50  c.c.  of  any  of  the 
chromo-acetic  solutions  is  often  an  advantage. 

Chromo- Acetic  Solution  (for  marine  algae).— 

Chromic  acid 1 . 0  g. 

Glacial  acetic  acid 0.4  c.c. 

Sea-water , 400. 0 c.c. 

Material  must  be  washed  in  sea-water. 

Flemming's  Fluid  (weaker  solution). — 

f  1  per  cent  chromic  acid  (in  water) 25  c.c. 

A  <  1  per  cent  glacial  acetic  acid  (in  water)  ....  10  c.c. 

I  Water 55  c.c. 

B.    1  per  cent  osmic  acid  (in  water) 10  c.c. 

Keep  the  mixture  A  made  up,  and  add  B  as  the  reagent  is  needed 
for  use,  since  it  does  not  keep  well. 

Flemming's  Fluid  (stronger  solution). — 

1  per  cent  chromic  acid 45  c.c. 

2  per  cent  osmic  acid 12  c.c. 

Glacial  acetic  acid 3  c.c. 

Merkel's  Fluid.— 

1.4  per  cent  solution  of  chromic  acid 25  c.c. 

1.4  per  cent  solution  of  platinic  chloride 25  c.c. 

Benda's  Fluid. — 

1  per  cent  chromic  acid 16  c.c. 

2  per  cent  osmic  acid 4  c.c. 

Glacial  acetic  acid 2  drops 

Hermann's  Fluid. — 

1  per  cent  platinic  chloride 15  parts 

Glacial  acetic  acid 1  part 

2  per  cent  osmic  acid 4  or  2  parts 


Formulae  for  Reagents  299 

Picric  Acid.— 

Picric  acid 1  g_ 

Water  or  70  per  cent  alcohol 100  c.c. 

Bouin's  Fluid. — 

Commercial  formalin 25  c.c. 

Picric  acid  (saturated  solution  in  water) 75  c.c. 

Glacial  acetic  acid 5  c.c. 

Corrosive  Sublimate  and  Acetic  Acid. — 

Corrosive  sublimate 3  g. 

Glacial  acetic  acid 3  c.c. 

70  per  cent  alcohol  (or  water) 100  c.c. 

Bensley's  Formula  (for  mitochondria). — 

2|  per  cent  corrosive  sublimate  in  water 4  parts 

2  per  cent  osmic  acid 1  part 

Corrosive  Sublimate,  Acetic  Acid,  and  Picric  Acid. — 

Corrosive  sublimate 5  g. 

Glacial  acetic  acid 5  c.c. 

Picric  acid 1  g. 

50  per  cent  alcohol 100  c.c. 

Corrosive  Sublimate  and  Picric  Acid  (Jeffrey's  formula). — 
Corrosive  sublimate,  saturated  solution  in  30 

per  cent  alcohol 3  parts 

Picric  acid,  saturated  solution  in  30  per  cent 

alcohol 1  part 

Gilson's  Fluid.— 

95  per  cent  alcohol 42  c.c. 

Water 00  c.c. 

Glacial  acetic  acid 18  c.c. 

Concentrated  nitric  acid 2  c.c. 

Corrosive    sublimate    (saturated    solution    in 

water) llc-c- 

Bensley's  Formula  (for  canal  system).— 

1.  Bichromate  of  potash 

2.  Corrosive  sublimate ^   g. 

3.  Water 90   c"c' 

4.  Formalin  (neutral) 

Make  a  solution  of  1,  2,  3,  and  then  add  the  neutral  formalin. 


300  Methods  in  Plant  Histology 

Osmic  Acid. — 

Osmic  acid 1  c.c. 

Distilled  water 100  c.c. 

The  bottle  in  which  the  solution  is  to  be  kept,  and  also  the  glass 
tube  in  which  the  acid  is  sold,  must  be  thoroughly  cleaned.  Break 
off  the  end  of  the  tube,  and  drop  both  tube  and  acid  into  the  distilled 
water,  or  simply  drop  the  tube  into  the  bottle  and  shake  the  bottle 

until  the  tube  breaks. 

STAINS 

Delafield's  Haematoxylin. — "To  100  c.c.  of  a  saturated  solution 
of  ammonia  alum  add,  drop  by  drop,  a  solution  of  1  g.  of  haematoxy- 
lin  dissolved  in  6  c.c.  of  absolute  alcohol.  Expose  to  air  and  light 
for  one  week.  Filter.  Add  25  c.c.  of  glycerin  and  25  c.c.  of  methyl 
alcohol.  Allow  to  stand  until  the  color  is  sufficiently  dark.  Filter 
and  keep  in  a  tightly  stoppered  bottle"  (Stirling  and  Lee). 

The  solution  should  stand  for  at  least  two  months  before  it  is 
ready  for  using. 

Erlich's  Haematoxylin. — 

Distilled  water 50  c.c. 

Absolute  alcohol 50  c.c. 

Glycerin 50  c.c. 

Glacial  acetic  acid 5  c.c. 

Haematoxylin 1  g. 

Alum  in  excess. 

Keep  it  in  a  dark  place  until  the  color  becomes  a  deep  red.  If 
well  stoppered,  it  will  keep  indefinitely. 

Boehmer's  Haematoxylin. — 

.    f  Haematoxylin 1  g. 

\  Absolute  alcohol 12  c.c. 

B   f  Alum 1  g. 

\  Distilled  water 240  c.c. 

The  solution  A  must  ripen  for  two  months.  When  wanted  for 
use,  add  about  10  drops  of  A  to  10  c.c.  of  B.  Stain  10  to  20  minutes. 
Wash  in  water  and  proceed  as  usual. 

Mayer's  Haem- Alum. — Haematoxylin,  1  g.,  dissolved  with  heat 
in  50  c.c.  of  95  per  cent  alcohol  and  added  to  a  solution  of  50  g.  of 


Formulae  for  Reagents  301 

alum  in  a  liter  of  distilled  water.     Allow  the  mixture  to  cool  and 
settle;   filter;   add  a  crystal  of  thymol  to  preserve  from  mold  (Lee). 
It  is  ready  for  use  as  soon  as  made  up.     Unless  attacked  by  mold, 
it  keeps  indefinitely. 

Haidenhain's  Iron-Haematoxylin. — This  stain  was  introduced 
by  Haidenhain  in  1892  and  has  gained  a  well-deserved  popularity 
with  those  engaged  in  cytological  work.  Two  solutions  are  used, 
and  they  are  never  mixed : 

A.  1^  to  4  per  cent  aqueous  solution  of  ammonia  sulphate  of  iron.     Use 
the  ferric  (violet)  crystals,  not  the  ferrous  (green)  crystals. 

B.  ^  per  cent  solution  of  haematoxylin  in  distilled  water. 

The  crystals  of  haematoxylin  will  dissolve  in  the  distilled  water 
in  about  10  days;  the  stain  reaches  its  greatest  efficiency  in  about 
6  weeks.  About  3  months  from  the  time  it  is  made  up,  it  begins  to 
deteriorate.  A  stain  made  by  dissolving  the  crystals  in  strong  alco- 
hol and  then  diluting  with  water  so  as  to  get  a  practically  aqueous 
solution  is  not  so  good. 

Greenacher's  Borax  Carmine. — 

Carmine 3  g. 

Borax 4  g. 

Distilled  water .100  c.c. 

Dissolve  the  borax  in  water  and  add  the  carmine,  which  is  quickly 
dissolved  with  the  aid  of  gentle  heat.  Add  100  c.c.  of  70  per  cent 
alcohol  and  filter  (Stirling) , 

Alum  Carmine.— A  4  per  cent  aqueous  solution  of  ammonia  alum 
is  boiled  20  minutes  with  1  per  cent  of  powdered  carmine.  Filter 
after  it  cools  (Lee). 

Alum  Cochineal.— 

Powdered  cochineal 

Alum 

Distilled  water 

Dissolve  the  alum  in  water,  add  the  cochineal,  and  boil;  evaporate 
down  to  two-thirds  of  the  original  volume  and  filter.  Add  a  few 
drops  of  carbolic  acid  to  prevent  mold  (Stirling). 


Jfeifcxb  «•  Plant  Histology 


Water,  or  70  per  cent  alcohol 100e-C- 

General  Formula  for  Anilms. — Make  a  3  per  cent  solution  of 
anilin  oil  in  distilled  water;  shake  well  and  frequently  for  a  day; 
add  enough  alcohol  to  make  the  whole  mixture  about  20  per  cent 
alcohol;  add  1  g-  of  cyaninT  erythroem.  safranin.  gentian-violet,  etc., 
to  each  100  c.c.  of  this  solution. 

Cyanin. — This  general  formula  is  not  at  all  successful  with 
Gruber s  eyanin.  but  gives  satisfactory  results  with  an  immensely 
cheaper  cyanin,  sold  by  H.  A.  Met*  &  Co.,  122  Hudson  Street,  New 
York. 

Aniltn  Blue. — 

Anilin  blue 1  g. 

So  or  90  per  cent  alcohol 100  e,c. 

For  staining  before  mounting  in  Venetian  turpentine,  this  stain 
should  be  made  up  in  strong  alcohol,  even  if  the  dry  stain  is  intended 
for  aqueous  solution. 

Iodine  Green. — 

Iodine  green 1  g. 

70  per  cent  alcohol 100  CJK. 

Methyl  Green.— 

Methyl  green 1  g. 

Glacial  acetic  acid 1  e-e. 

Water 100e.e. 

If  the  preparation  is  to  be  mounted  in  balsam,  a  slight  trace  of 
acetic  acid  and  also  a  trace  of  methyl  green  should  be  added  to  the 
absolute  alcohol  used  for  dehydrating. 

For  staining  vascular  bundles,  the  acid  may  be  omitted,  even  from 
the  formula. 


Formulae  for  Reagent*  303 

Light  Green.— 

Light  gran.  ...  lg 


Clove  oil  100c_0_ 


r 


Light  green j  c 

Cloveoil 75o.o. 

Absolute  alcohol 25  o.o. 

Fuchsin. — 

Fuehsin 1  g 

95  per  cent  alcohol 100  o.o. 

Water...                          100  o.o. 

Acid  Fuchsin. — 

Acid  fuchsia \  jr. 

Water.  .                         100  c.o. 

Use  this  formula  when  staining  woody  tissues  in  methyl  green 
and  aeid  fuehsin. 

Ziehl's  Carbol  Fuchsin.— 

Fuohsin 1  g. 

Carbolic-acid  crystals 5  g. 

95  per  cent  alcohol 10  o.o. 

Water 100  c.o. 

Fuchsin  and  Iodine  Green  Mixtures. — Two  solutions  are  kept 
separate,  since  they  do  not  retain  their  efficiency  long  after  they  an" 
mixed. 

Fuchsin  (acid) 0. 1  g. 

\  Distilled  water 50. 0  c.o. 

I  inline  green 0. 1  g. 

13  \Pistillod  water 50.0  c.o. 

I  Absolute  alcohol 100. 0  c.o. 

C  <j  Glacial  acetic  acid 10  c.o. 

.Iodine 0.lR. 

Stain  in  equal  parts  of  A  and  B.  Transfer  from  the  stain  directly 
to  solution  C,  and  from  C  to  xylol. 


304  Methods  in  Plant  Histology 

Another  Formula. — 

f  Acid  fuchsin 0 . 5  g. 

A  \  Water 100.0  c.c. 

J  Iodine  green 0 . 5  g. 

B  \Water 100.0 c.c. 

Mix  a  pipette  full  of  A  with  a  pipette  full  of  B ;  stain  2  to  8  min- 
utes; dehydrate  rapidly  and  mount  in  balsam. 

Magdala  Red.— 

Magdala  red 1  g. 

85  or  90  per  cent  alcohol 100  c.c. 

Use  this  formula  when  staining  in  Magdala  red  and  anilin  blue, 
before  mounting  in  Venetian  turpentine. 

Safranin. — 

Safranin 1  g. 

95  per  cent  alcohol 50  c.c. 

Water 50  c.c.    . 

Safranin  (another  formula) .-  -  • 

Dissolve  1  g.  of  alcohol-soluble  safranin  in  100  c.c.  of  absolute  alcohol; 
dissolve  1  g.  of  water-soluble  safranin  in  100  c.c.  of  distilled  water.  Mix 
equal  parts  of  the  two  solutions. 

Gentian-Violet. — 

Gentian-violet 1  g. 

95  per  cent  alcohol 20  c.c. 

Water. 80  c.c. 

Anilin  oil 3  c.c. 

A  1  per  cent  solution  in  water  keeps  better. 

A  1  per  cent  solution  in  clove  oil  is  worth  a  thorough  trial. 

Pyoktanin. — This  is  sold  by  E.  Merck,  in  Darmstadt,  Germany. 
Dissolve  1  g.  of  pyoktanin  in  30  c.c.  of  water. 

Orange  G. — 

Orange  G 1  g. 

Water 100  c.c. 

For  most  purposes  a  1  per  cent  solution  in  clove  oil  is  preferable. 


Formulae  for  Reagents 


Gold  Orange.— 

Gold  orange 

Clove  oil 

Bismarck  Brown. — 

Bismarck  brown 
70  per  cent  alcohol . 


100  c.c. 


2g. 

100  c.c. 


Nigrosin.— 

Nigrosin 1  g 

Water ,. 100  c.c. 

Gram's  Solution.— 

Iodine. j  g 

Iodide  of  potassium 2  g. 

Water 300  c.c. 

MISCELLANEOUS 

Mayer's  Albumen  Fixative. — 

White  of  egg  (active  principle) 50  c.c. 

Glycerin  (to  keep  it  from  drying  up) 50  c.c. 

Salycilate  of  soda  (to  keep  out  bacteria,  etc.) . .       1  g. 
Shake  well  and  filter. 

Land's  Gum  Fixative. — 

Gum  arabic 1  g. 

Potassium  bichromate 1  g. 

Water 98  c.c. 

Dissolve  the  gum  in  water  and  add  the  bichromate  of  potash; 
or  dissolve  the  gum  in  half  the  quantity  of  water  and  the  bichromate 
of  potash  in  the  other  half,  and  mix  just  before  using.  Le  Page's 
liquid  glue  may  be  used  instead  of  the  gum  arabic. 

Schultze's  Maceration  Fluid. — 

The  ingredients  are  nitric  acid  and  potassium  chlorate.  They  are 
mixed  only  as  the  reagent  is  applied.  See  chapter  on  "Special  Methods" 
(chap.  xi). 

Iodine  (solution  for  starch  test). — 

Dissolve  1  g.  potassium  iodide  in  100  c.c.  of  water  and  add  0.3  g.  sub- 
limed iodine. 


306  Methods  in  Plant  Histology 

Fehling's  Solution. — 

f  Cupric  sulphate 3  g. 

A  \  Water 100  c.c. 

j  Sodium  potassium  tartrate  (Rochelle  salts)  16  g. 

B  (  Water 100  c.c. 

f  Caustic  soda 12  g. 

U  \  Water' 100  c.c. 

Keep  it  in  three  bottles  labeled  A,  B,  and  C.  When  needed  for 
use,  add  10  c.c.  of  water  to  5  c.c.  from  each  of  the  three  bottles. 

Millon's  Reagent. — 

Mercury 1  c.c. 

Concentrated  nitric  acid 9  c.c. 

Water '. 10  c.c.. 

Cuprammonia. — 

Prepare  by  pouring  15  per  cent  ammonia  water  upon  copper  turnings 
or  filings.  Let  it  stand  in  an  open  bottle. 

Phloroglucin . — 

Use  a  5  per  cent  solution  in  water  or  alcohol. 

Celloidin.— 

To  make  a  2  per  cent  solution,  add  one  tablet  of  Schering's  celloidin  and 
enough  ether-alcohol  (equal  parts  absolute  alcohol  and  ether)  to  make  the 
whole  weigh  2,000  g. 

Where  only  a  small  quantity  is  needed,  shave  off  2  g.  of  celloidin  and  add 
100  c.c.  of  ether  alcohol. 

Eycleshymer's  Clearing  Fluid.— 

Mix  equal  parts  of  bergamot  oil,  cedar  oil,  and  carbolic  acid. 

Venetian  Turpentine. — 

To  make  a  10  per  cent  solution,  add  90  c.c.  of  absolute  alcohol  to  10  c.c. 
of  thick  Venetian  turpentine.  Stir  it  with  a  glass  rod.  Guess  at  the  amount 
of  turpentine,  for  it  is  not  easy  to  clean  things  which  have  contained  Venetian 
turpentine. 

The  following  need  no  formulae:  Acetic  acid,  hydrochloric  acid, 
nitric  acid,  sulphuric  acid,  carbolic  acid,  chloroform,  ether,  xylol, 
cedar  oil,  clove  oil,  bergamot  oil,  turpentine,  glycerin,  paraffin, 
balsam. 


Formulae  for  Reagents  307 

AMOUNTS  OF  REAGENTS  REQUIRED 

It  is  difficult  to  estimate  the  amounts  of  the  various  reagents 
needed  by  a  class  in  histology.  Two  dangers  must  be  guarded 
against — wastefulness  and  too  great  economy;  for  economy  in  some 
reagents,  like  absolute  alcohol,  turpentine,  xylol,  and  clove  oil,  may 
be  so  rigid  as  to  make  the  preparations  decidedly  inferior. 

Each  student  should  have  some  reagents  upon  his  own  table. 
The  following  is  an  estimate  of  the  amounts  of  some  reagents  used 
by  each  student  in  a  three-months'  course  in  method:  commercial 
alcohol  (about  95  per  cent),  3  liters;  absolute  alcohol,  800  c.c.; 
turpentine  (for  dissolving  paraffin  ribbons),  200  c.c.;  xylol,  400  c.c.; 
clove  oil,  75c.c.;  Canada  balsam,  25  c.c.;  hard  and  soft  paraffin, 
400  g.  each;  safranin,  gentian-violet,  orange,  cyanin,  erythrosin, 
Delafield's  haematoxylin,  iron-haematoxylin,  and  ammonia  sulphate 
of  iron  (3  per  cent),  100  c.c.  each. 

For  general  use  of  the  entire  class,  other  stains  and  reagents  may 
be  kept  upon  a  table  accessible  to  all.  Some  stains  which  act  very 
rapidly,  like  cyanin,  erythrosin,  and  orange,  may  be  kept  upon  the 
common  table.  A  class  of  ten  will  use  about  20  liters  of  the  stock 
solution  of  chromo-acetic  acid,  and  of  glacial  acetic  acid  about 
400  c.c.;  commercial  formalin,  about  200  c.c.;  Venetian  turpentine, 
500  c.c. ;  cedar  oil,  200  c.c.;  Eycleshymer's  clearing  fluid,  100  c.c.; 
glycerin,  600  c.c.;  chloroform,  100  c.c.;  ether-alcohol,  200  c.c.; 
celloidin,  40  g.;  hydrochloric  acid,  200  c.c.;  nitric  acid  and  sulphuric 
acid,  50  c.c.  each;  Magdala  red  and  methyl  green,  200  c.c.  each; 
other  stains,  100  c.c.  each. 

No  attempt  has  been  made  to  make  the  foregoing  list  absolutely 
complete.    The  equipment  of  any  laboratory  will  be  built  up  gradu- 
ally.    When  the  microtome  needs  oiling,  some  sewing-machine  c 
may  be  added  to  the  general  equipment.     It  is  hoped  that  reagents 
omitted  from  the  list  are,  like  the  sewing-machine  oil,  readily  sc 
without  vexatious  delays. 


INDEX 


INDEX 


[The  references  are  to  pages.     Italic  figures  indicate  illustrations.] 


Abies,  256 
Acetic  alcohol,  19. 
Achlya,  195. 
Acid  fuchsin,  53. 
Acid  green,  58. 
Acorus,  262. 
Aecidium,  204. 
Agaricus,  205. 
Agathis,  255. 
Albugo,  195. 
Alcohol,  18. 
Allium,  263. 
Anabaena,  160. 
Anemone,  276. 
Aneura,  213. 
Angiopteris,  236. 
Angiosperms,  261. 
Anilin,  50. 

Anilin  blue,  58,  62,  100. 
Anthers,  fixing,  102. 
Anthoceros,  214. 

Asclepias,  pollen,  273. 
Asperaillus,  19S. 
Asterella,  210. 
Azolla,  240,  243. 

Bacteria,  154. 
Balsam,  Canada,  37.    . 
Bath,  12,  14,  108. 
Batrachospermum,  188,  189. 
Benda's  fluid,  26. 
Bensley's  formula,  30. 
Bergamot  oil,  34. 
Bismarck  brown,  58. 
Blue,  anilin,  58,  62,  100. 

Boletus  194. 

Botrychium,  230,  SSI. 
Bourn's  fluid,  27. 

Bovista,  206. 

Bryophytes,  207. 

Bryum,  219,  220. 

Buds,  265,  267. 

Callithamnion,  190. 

Callose,  77. 

Canaliculi,  133,  134. 

Cane-sugar,  75. 

Capsella,  266,  278. 

Carbolic  acid,  34. 


Carmine,  Greenacher's  borax,  49. 

Carnoy's  fluid,  19. 

Cataloging,  288. 

Cedar  oil,  35. 

Celloidin  method,  119. 

Cellulose,  76. 

Ceratozamia,  104,  S46,  S48. 

Chara,  180. 

Chloroform,  35. 

Chlorophyceae,  162. 

Chromic  acid,  20. 

Cilia,  131. 

Circium,  floral  development,  267. 

Cladophora,  163,  179. 

Clearing,  34,  106,  116. 

Clintonia,  Stem,  262. 

Closterium,  164. 

Clove  oil,  35. 

Coelosphaerium,  160. 

Collema,  202. 

Combination  stains,  60. 

Congo,  red,  53,  87. 

Conocephalus,  210. 

Coplin  jar,  1">. 

Coprinus,  205,  206. 

Corallina,  190. 

Corrosive  sublimate,  28. 

Covers,  16. 

Crucibulum,  206. 

Crystals,  78. 

Cultures  of  algae,  163. 

Cutin,  78. 

Cutleria,  182,  184. 

Cutting,  111. 

Cyanin,  56. 

Cyanin  and  erythrosin,  61. 

Cyanophyceae,  156. 

Cyathus,  206. 

Cycadales,  244. 

Cypripedium,  275. 

Dehydration,  31,  105,  116,123. 
Desifllication,  123. 
Desmids,  173. 
Desmotrichum,  182,  183. 
Developer,  Land's,  143. 
Diatoms,  171,  171. 
Dictyota,  182,  187. 
Dioon,  245. 


311 


312 


Methods  in  Plant  Histology 


Ectocarpus,  182,  183. 

Embryo:  angiosperms,  279;     Pinus,  260. 

Embryo-sac,  276. 

Endosperm,  277. 

Eosin  54. 

Epidermis,  91,  266. 

Equisetales,  227. 

Equisetum,  228. 

Erysiphe,  199. 

Erysipheae,  199. 

Erythronium,  pollen,  271. 

Erythrosin,  54. 

Eudorina,  166. 

Euglena,  166. 

Eurotium,  197. 

Fats,  76. 

Fertilization,  277. 

Filicales,  233. 

Fixative:  Land's,  114;   Mayer's,  113. 

Fixing  agents,  17. 

Fixing,  general  hints,  30. 

Flemming's  fluid,  25. 

Flemming's  triple  stain,  59. 

Floral  development,  266. 

Forceps,  14. 

Formalin,  29. 

Formalin  alcohol,  20. 

Formulae,  297. 

Freehand  sections,  80. 

Fuchsin,  acid,  53. 

Fucus,  185,  186. 

Funaria,  216,  218,  219. 

Fungi,  192. 

Geaster,  206. 

Gelatin,  78. 

Gelatin  method,  Land's,  128. 

Gentian-violet,  55 

Gilson's  fluid,  205. 

Ginkgoales,  250. 

Gleichenia,  235. 

Gloeotrichia,  159. 

Glycerin,  37. 

Glycerin  method,  92. 

Gold  orange,  58. 

Gonium,  166. 

Graduate,  15. 

Grape-sugar,  75. 

Green:  acid,  58;    iodine,    57;    light,    57; 

malachite,  57;    methyl,  57. 
Green  tones,  lantern  slides,  146. 
Griffithsia,  190. 
Gum,  78. 

Gymnosperms,  244. 
Gymnosporangium,  205. 

Haematoxylin,  40;   Delafleld's,  45. 
Hanging-drop  cultures,  73. 


Hardening,  105. 
Helminthostachys,  230. 
Hemiascomycetes,  196. 
Hepaticae,  207. 
Herbaceous  sections,  81. 
Hones,  11. 
Hydnum,  206. 
Hydrodictyon,  168. 
Hypoxylon,  201. 

Imbedding:    in  celloidin,  124;    in  gelatin, 

129;  in  paraffin,  109. 
Infiltration:     with    celloidin,    124;     with 

gelatin,  128;  with  paraffin,  107. 
Iodine,  28. 
Iodine  green,  57. 
Iris,  embryo,  279. 

Iron-alum  haematoxylin,  Haidenhain's,  41. 
Isoetes,  225. 

Juniperus,  255. 

Killing  agents,  17. 
Klebs's  culture  methods,  163. 
Knife,  position,  112. 
Knop's  solution,  164. 

Labeling,  84,  288. 

Laminaria,  182,  183. 

Lantern    slides,    141;     intensifying,    144; 

reducing,  144;   toning,  145. 
Large  sections,  125. 
Leaf,  264. 

Leaves  of  cycads,  246. 
Leaves  of  Pinus,  253. 
Light  green,  57. 
Lignin,  77. 
Lilium,  109;    archesporium,  872;   oogen- 

esis,   274;   ovary,    273;    pollen   mother 

cells,  269. 
Liverworts,  90. 
Living  tissues,  135. 
Lycoperdon,  206. 
Lycopodiales,  221. 
Lycopodium,  222. 

Maceration  method,  Schultze's,  129. 
Magdala  red,  54;  with  anilin  blue,  62,  100. 
Malachite  green,  57;   with  Congo  red,  87. 
Marchantia,  208,    214;     antheridia,    210; 

archegonia,  211. 
Marine  forms  of  algae,  165. 
Marsilia,  240,  242. 
Methyl  green,  57. 
Microchemical  tests,  74. 
Micrometry,  280. 
Micron,  282. 
Microscope,  5,  6,  281. 
Microtome,  7,  9. 


Index 


313 


Microtome  knives,  9. 

Microtome  with  motor,  los. 

Middle  lamella,  70. 

Mitochondria,  132,  1,3.3. 

Mitosis:    pollen   mother  cells,    268,    269; 

root-tips,  262,  263. 
Moonlight  tints,  lantern  slides,  146. 
Mosses,  90. 

Mounting  in  balsam,  117. 
Mucilage,  78. 
Mucor,  192. 
Musci,  216. 
Myxomycetes,  152. 

Nemalion,  188,  189. 
Needles,  13. 
Xidularia,  206. 
Xigrosin,  59 
Nostoc,  158. 
Nummularia,  201. 

Oedogonium,  163,  170. 

Oils,  76. 

Oogenesis:      angiosperms,     274;      gymno- 

sperms,   248,    255,    257;    pteridophytes 

240,  241. 

Ophioglossum,  230,  232. 
Orange  G,  58. 
Oscillatoria,  156,  157. 
Osmunda,  234,  236,  241. 

Pandorina,  166. 

Paraffin,  37;    cakes,  no;   method,  102. 

Parmelia,  202. 

Pellia,  209,  210,  21J. 

Peltigera,  202. 

Penicillium,  199. 

Peristome,  90. 

Petrifactions,  126. 

Peziza,  196,  197. 

Phaeophyceae,  181. 

Photomicrographs,  136. 

Phycomycetes,  192. 

Physcia,  202. 

Phytelephas,  131. 

Picric  acid,  27. 

Pilularia,  240. 

Pinus,  109,  251,  257,  258,  259. 

Plasmodia,  153. 

Pleurococcus,  166. 

Podocarpus,  255. 

Pollen  grain,  271,272. 

Polyporus,  206. 

Polysiphonia,  190,  191. 

Polytrichum,  217. 

Proteids,  75. 

Prothallia,  21,  89,    101;    Equisetum,  S28; 

Filicales,  237,  2-33,  240,  241;     Marsilia, 

£42;    Selaginella,  235. 


Protonema,  216. 
Protoplasmic  connections,  129. 
Prunus,  floral  development,  267. 
Pteridophytes,  221,  227,  230.  233 
Pteris,  233,  234,  238. 
Ptilidium,  208. 
Puccinia,  202,  203. 

Quercus,  261. 

Ranunculus,  266;   floral  development,  266; 

root,  264. 

Reduction  of  chromosomes,  275. 
Ribbons,  112. 
Riccia,  207,  212. 
Rii-ularia,  158. 
Rhizopus  (Mucor),  192,  133. 
Rhodophyceae,  188. 
Roots,  262;    Pinus,  253;    tips,  102,  262. 

Saccharomycetes,  196. 
Safranin,  51;    with  anilin  blue,  85;    with 
haematoxylin,  81;    with  light  green,  86. 
Sagittaria,  embryo,  279. 
Salix,  floral  development,  267. 
Salvinia,  240. 
Saprolegnia,  194,  195. 
Scalpels,  10. 
Scenedesmus,  166,  167. 
Schedules:    glycerin,    94;    celloidin,    122; 
freehand     sections,     84;      iron-haema- 
toxylin,  98;    paraffin,  117;    Sharp's,  43; 
Venetian  turpentine,  98;   Yamanouchi's. 
42. 

Scleroderma,  206. 
Sealing,  94. 
Sedum,  266. 
Selaginella,  223,  22.',. 
Senecio,  275. 

Sepia  tones,  lantern  slides,  145. 
Sieve  tubes,  262. 
Siphonostele,  261. 
Slides,  15. 
Sori,  fern,  90. 
Sparganium  root,  26 5. 
Spermatogenesis:    anKiosperms,  208,  tH9; 
gymnosperms,    246,    24S,  253;  pterido- 
phytes, 228,  238. 
Spermatophytes,  244. 
Sphacelaria,  182. 
Sphaerotheca,  199. 
Sphagnum,  220. 
Sphenophyllales,  221. 
Spirogyra,  163,  164,  /  75. 
Sporodinia,  194. 
Sporogenesis.  236,  237. 
Staining-dishes.  15.  40. 
Stains,  combination,  59. 


314 


Methods  in  Plant  Histology 


Starch.  74. 

Stem:     angiosperm,    261;      Pinus, 

pteridophyte,  230,  234,  235. 
Stender  dish,  15. 
Sticta,  202. 
Stony  tissues,  126. 
Strickler's  clamp,  10. 
Stypocaulon,  182. 
Suberin,  78. 
Synthol,  33. 

Taraxacum,  floral  development,  267. 

Taxodium,  255. 

Temporary  moimts,  72. 

Telracera,  262. 

Thermostat,  12,  13. 

Thick  sections,  128. 

Thuja,  255. 

Tilia,  261. 

Tolypothrix,  168. 

Tradescanlia,  root-tip,  £62. 

Transfer  to  paraffin,  107. 

Trillium,  275. 

Tube  length,  282. 


Turntable,  12,  14. 

Typha,  floral  development,  267. 

Ulolhrix,  163,  169. 
Uncinulc,  199,  200. 
Uredineac,  203. 
Usnea,  202. 
Ustilagineae,  202. 
Usta-l-.ia,  201. 

Vaucheric,  177. 
Venetian  turpentine,  97. 
Ftcto  Faba,  263. 
Volvox,  165,  166. 

Washinc,  23,  104. 
Wr^scrbluthe,  160. 

Woody  sections,  81. 

Xylol,  34. 
Xylaria,  201. 

Zamia,  244,  24». 

Zea  Mays:  anatomy,  264;  embryo,  279. 

Zygnema,  174. 


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